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DISSERTATION

Titel der Dissertation

Community structure and distribution of functional microbial groups within two complex environments: associated with marine and potential sulfur-compounds-reducing microorganisms in peatlands.

angestrebter akademischer Grad

Doktorin der Naturwissenschaften (Dr. rer.nat.)

Verfasserin / Verfasser: Doris Steger Matrikel-Nummer: 9803561 Dissertationsgebiet (lt. Stu- 444 Ökologie dienblatt): Betreuerin / Betreuer: Univ.-Prof. Dr. Michael Wagner

Wien, am 18 März 2010

Formular Nr.: A.04

Contents

Outline ......1

Part I Sulfate reducers in peatlands

Introduction 5

Microorganisms with Novel Dissimilatory (Bi)Sulfite Reductase Genes are Widespread in Geographically Distant, LowSulfate Peatlands .21

Part II Hybridization regimes in DNA microarray experiments

Introduction .....79

Spatial Gradients of Target Nucleic Acids Govern DNA Microarray Hybridization Assays ..87

Part III Microorganisms in marine Sponges

Introduction 113

SpongeAssociated Microorganisms: , Ecology, and Biotechnological Potential .125

Diversity and Mode of Transmission of Ammoniaoxidizing in Marine Sponges .195

Part IV Summary ....205

Appendix ..211

Outline

Part I of this thesis describes the analysis of dsrABcarrying microorganisms in terrestrial wetlands. Newly developed tools such as a functional gene microarray, quantitative PCR assays and dsrB based denaturing gradient gel electrophoresis as well as clone libraries were applied to unravel the widespread occurrence of deepbranching dsrAB lineages, which outnumbered characterized dsrAB lineages in the Schlöppnerbrunnen fen system.

Part II validates the signal variability in DNA microarray hybridizations. A simplified microarray format and numerical simulation tests were applied to investigate systematic variations in detected signal intensities and to explore underlying mechanisms of observed spatial gradients.

Part III gives a comprehensive overview on the community structure of associated microorganisms. A metaanalysis of all public available sponge derived 16S rRNA gene sequences as well as of new sequences retrieved from so far unstudied marine sponges was conducted. Moreover, the mode of transmission of Archaea probably involved in ammonia oxidation was analyzed by 16S rRNA and ammonia monooxygenase gene based clone libraries. The presence of detected putative ammoniaoxidizing archaea (AOA) in adult sponges and corresponding sponge larvae was confirmed by fluorescence in situ hybridization.

Part IV includes a summary of the findings obtained in this thesis

The Appendix includes a list of the author’s publications, oral and poster presentations, the acknowledgment, and the curriculum vitae.

1

Part I

Sulfate reducers in peatlands

Sulfate reducers in peatlands

Introduction

1. Peatlands and their role in the global carbon cycle

Wetlands are a common part of the global landscape, with mainly three distinguishing characteristics to other landform units: (i) the permanent or periodic presence of a table at, near or above the land surface, which determines (ii) a unique development that is frequently characterized by low content and (iii) specialized communities that are adapted to these environments (Cowardin et al. 1995). Peatlands, also called organic wetlands, build up a substantial fraction of these systems and are probably the most widespread type of wetlands by occupying globally an area of about 4*106 km2 (Joosten et al. 2002). They represent ecosystems where net exceeded the rate of decomposition and as a result predominately organic, carbon rich matter, so-called peat, accumulated over time. Their persistence depends on a constant long-term water input, and the origin of this water influences the form and function of peatlands. Based on their main water source, peatlands can be divided in predominantly groundwater-fed fens (minerotrophic peatlands) and in rainwater-fed bogs (ombrotrophic peatlands) (Bridgham et al. 1996). The source of water considerably influences the geochemistry of the peatland. Minerotrophic fens receive nutrient-input from incoming mineralized groundwater and thus higher concentrations of cations and anions are present compared to exclusively precipitation-fed bogs. Bogs are oligotrophic, as isolation from surrounding mineral soil results in low nutrient and mineral content, which is often associated with low pH values (< 5) (Bridgham et al. 1996). Dominating vegetation forms are Sphagnum mosses, sedges and ericaceous shrubs, which are adapted to these conditions (Vanbreemen 1995).

Peatlands store about 15–30% of the world’s soil carbon (Turunen et al. 2002; Davidson et al. 2006). Thus, they are massive deposits of carbon, acting as a net carbon sink and contributing to global cooling in the past 8,000 to 11,000 (Frolking et al. 2007). Undisturbed peatlands are likely to continue functioning as net carbon sinks at the decadal scale, despite the large interannual variability of individual peatlands (Moore et al. 1998). However, environmental and anthropogenic factors, such as climate change and land-use, affect the carbon balance of these ecosystems and can lead to substantial carbon loss (Dise 2009). Primary carbon inputs in

5 Part I peatlands derive from plantassimilated carbon dioxide, whereby energyrich organic compounds are deposited as litter or released belowground by plant roots. Carbon loss is dominated by the emission of the gases carbon dioxide and methane (Keller et al. 2007).

complex polymers

exoenzymes denitrifiers - NO3 → N2 monomers manganese reducers Mn4+ → Mn2+

iron reducers

redox potential primary fermenters Fe3+ → Fe2+ Acidiphilium-, Geobacter-, Clostridiaceae, , and Geothrix-related spp. Actinomycetales,

sulfate reducers 2+ 2- SO4 → S Desulfomonile-, Desulforhopalus- and secondary fermenters acetogens Desulfosporosinus–related spp. Syntrophobacter-related spp. Clostridiaceae, Acidaminococcaceae methanogens

CO2 → CH4 Methanobacterium-, Methanosaeta-, Methanosarcina- and Methanospirillum-related spp.

Figure 1: Carbon mineralization processes in peatlands. Extracellular enzymes hydrolyze complex to monomers. Organic substrates, methanol or carbon dioxide are oxidized to acetate by acetogens or degraded by primary fermenters to organic acids, alcohols and amino acids. Resulting products are available either for secondary (syntrophic microorganisms) or for the microbial mediated sequential reduction of nitrate, manganese, iron (III), sulfate and finally carbon dioxide. Microorganisms identified in the Schlöppnerbrunnen fen are displayed.

Soil microorganisms that use either oxygen or other electron acceptors as energy sources to decompose organic matter to carbon dioxide, methane and dissolved organic carbon play a major role in carbon effluxes from peatlands. As oxygen supply in peatlands is commonly restricted to small volumes, mainly anaerobic microbial mineralization using alternative electron acceptors takes place (Figure 1). Crucial factors influencing these decomposition processes are on one hand the availability of electron acceptors and on the other hand nutrient content and degradability of organic matter. In this context, the activity and potential environment-mediated inhibition of extracellular enzymes play a key role for substrate supply and microbial growth, as they hydrolyze complex organic matter to low molecular weight compounds (Makoi et al. 2008). Subsequently, acetogens are capable to oxidize a range of organic substrates (e.g. ),

6 Sulfate reducers in peatlands methanol or carbon dioxide with hydrogen as electron donor to produce acetate. Furthermore, primary fermenters degrade sugars to fermentation products, such as formate, lactate, butyrate and propionate. Resulting end products of these transformations are then available either for secondary fermentation catalyzed by syntrophic microorganisms or for several microbially mediated anaerobic reduction processes. Generally, a sequential reduction of nitrate, manganese, iron (III), sulfate and finally carbon dioxide is assumed, whereby respective respiration processes are regulated by differences in energy yields of the reactions (Achtnich et al. 1995). However, several studies have shown that respective processes can occur spatially and temporally at the same time in these heterogeneous and highly structured systems (Wieder et al. 1990; Yavitt et al. 1990; Koretsky et al. 200). Processes using inorganic electron acceptors often do not fully explain total anaerobic mineralized carbon amounts in peatlands, and thus the presence of other organic electron acceptors, such as humic acids, and fermentation processes coupled to methanogenesis are supposed to contribute substantially to the decomposition of organic compounds (Lovley et al. 1996; Vile et al. 2003; Keller et al. 200; Wüst et al. 2009).

2. Biogeochemical cycling of sulfur compounds in peatlands

Microbial mediated transformations of sulfur compounds are closely linked to the carbon cycle as reducing processes are associated with organic matter utilization in anoxic habitats such as peatlands. Peatlands have water saturated anoxic and overlying oxic zones with fluctuating water tables, and thus dynamic sulfur cycling is expected (Figure 2).

7 Part I

2 O SO4 2 H2S DMS

2 atmospheric SO4 deposition

su rf ac e inf low O2

2 2 oxic R―SO4 SO4 H2S 2- SO4 assimilation Absorption on DOM minerals CBS 2 2 S2O3 SO4 g rou ndw ate r anoxic flow 2 2 SO4 H2S SO4

Fe3+ RIS

FeS2 ,FeS DMS

2 Figure 2: Biogeochemical sulfur cycle in peatlands. Sulfate (SO4 ) can be assimilated, 2 2 adsorbed on minerals or forms organic sulfate esters (RSO4 ). Microbial dissimilatory SO4 reduction results in (H2S) production. In oxic zones, H2S gets reoxidized chemically to sulfate. H2S can also react to organic or carbon bonded sulfur (CBS) or inorganic reduced sulfur (RIS). RIS and CBS pools can be re-oxidized either aerobically or anaerobically. Chemical oxidation of H2S by dissolved organic matter (DOM) to thiosulfate, which subsequently can be utilized by microorganisms for thiosulfate reduction or disproportionation 2- into H2S and SO4 are proposed. Microbial mediated dimethyl sulfide (DMS) production in peatlands is suggested, which would be available for microbial reduction. Exclusively microbial driven transformations are indicated with dotted lines.

2- In bogs, sulfur is mainly introduced as sulfate (SO4 ) by atmospheric deposition, while in fens surface-water and groundwater inflows can contribute considerably to the sulfate pools. Sulfate can be assimilated by and microorganisms, adsorbed on soil minerals or forms organic 2- sulfate esters (R-SO4 ), which may hydrolyze under sulfate-poor conditions (Mandernack et al. 2000; Clark et al. 2005). In the anoxic zone, sulfate-reducing microorganisms (SRM) use sulfate to gain energy via dissimilatory sulfate reduction resulting in the production of sulfide. Sulfide reacts with hydrogen to hydrogen sulfide (H2S), which diffuses into the atmosphere or gets re- oxidized chemically to sulfate in oxic zones (Clark et al. 2005). H2S can also react with organic matter to form organic or carbon bonded sulfur (CBS) (Wieder et al. 1988) or with inorganic compounds, such as iron, contributing to the inorganic reduced sulfur (RIS) pool (Chapman 2001). RIS and CBS pools tend to be unstable in peatlands and can be re-oxidized either aerobically with oxygen if the water table drops (Mandernack et al. 2000), or anaerobically,

8 Sulfate reducers in peatlands using iron(III) or other compounds as anaerobic electron acceptor (Wieder et al. 1988; Blodau et al. 200). An alternative pathway could be chemical oxidation of H2S by dissolved organic matter to thiosulfate (Heitmann et al. 2006; Heitmann et al. 200), for example by quinone moieties of humic acids (Blodau et al. 200). Subsequently, thiosulfate can be utilized by microorganisms for thiosulfate reduction to H2S or oxidation to sulfate, respectively, or might be disproportionated into H2S and sulfate and may thus maintain the activity of present SRM (Jorgensen 1990). Finally, microbial mediated dimethyl sulfide production in peatlands is suggested (Kiene et al. 1995). Nevertheless, emissions of dimethyl sulfide from peatlands may be insignificant as it seems to be degraded rapidly by SRM and methanogens in freshwater sediments (Lomans et al. 1999).

3. The guild of sulfate-reducing microorganisms and their diversity in wetland systems

SRM represent a phylogenetically and metabolically diverse functional guild of microorganisms. Based on phylogenetic analysis of their 16S rRNA genes, at present, recognized SRM are affiliated to seven different phylogenetic phyla, two within the Archaea (i.e. the euryarchaeotal genus Archaeoglobus (Stetter 19) and the crenarchaeotal genera Caldivirga (Itoh et al. 1999) and Thermocladium (Itoh et al. 199)), and five within the . The major fraction of sulfatereducing bacteria belongs to the class Deltaproteobacteria, followed by the Firmicutes harboring the sulfatereducing genera Desulfotomaculum (Skerman et al. 190), Desulfosporosinus (Stackebrandt et al. 199), and Desulfosporomusa (Sass et al. 2004). Additionally, thermophilic SRM affiliated to Nitrospira (Thermodesulfovibrio spp. (Henry et al. 1994)), Thermodesulfobacteria (i.e. the genera Thermodesulfobacterium (Zeikus et al. 193) and Thermodesulfatator (Moussard et al. 2004)) and the family Thermodesulfobiaceae (Thermodesulfobium narugense (Mori et al. 2003)) were isolated from various environments.

SRM are characterized by their capability to gain energy for growth and by coupling the oxidation of organic compounds or hydrogen to the reduction of sulfate to sulfide in a process termed dissimilatory sulfate reduction (Rabus et al. 2000). The multistep process of dissimilatory sulfate reduction takes place in the cytoplasm or in association with the inner side of the cytoplasmic membrane and involves the transfer of eight electrons. Before being

9 Part I reduced, sulfate has to be activated with adenosine5’phosphate (ATP) by an ATP sulfurylase resulting in adenosine5’phosphosulfate (APS) and pyrophosphate. Subsequently, APS is converted to (bi) sulfite by an APS reductase, thereby two electrons are transferred to the sulfur ion and adenosine5’monophosphate is released. The reduction of sulfite to sulfide involves the transfer of 6 electrons and is catalyzed by a dissimilatory (bi)sulfite reductase (DSR). The electron transport reactions lead to the formation of a proton motive force that drives ATP synthesis by an ATPase (Rabus et al. 2000). Many SRM are capable to grow also by using numerous other electron acceptors, such as sulfite, thiosulfate, dimethylsulfoxide, nitrate or elemental sulfur (Dalsgaard et al. 1994; Moura et al. 199; Sass et al. 2009), which are reduced to sulfide as well. Based on their capability to oxidize lowmolecular mass organic compounds either completely or incompletely, two main physiological groups are roughly distinguished: SRM involved in incomplete oxidation of organic compounds resulting in acetate as an end product, or those capable of complete oxidation resulting in carbon dioxide (Muyzer et al. 200). Thereby, dissimilatory sulfate reduction leads to a relatively low energy gain, as for example the free energy change (∆G) of complete oxidation of acetate or lactate is 4 or 128 kJ/mol, respectively (Rabus et al. 2000). In the absence of sulfate, several SRM can also switch to the fermentation of organic substrates, or shift to a syntrophic lifestyle, such as Syntrophobacter (de Bok et al. 2002).

Several studies used 16S rRNA gene based approaches to investigate SRM diversity and distribution in complex systems. For instance, denaturing gradient gel electrophoresis targeting the 16S rRNA genes were used to detect selected groups of SRM in marine water columns and cyanobacterial mats (Teske et al. 1996; Teske et al. 1998), in industrial bioreactors (Dar et al. 2005) or in industrial paper mill (Maukonen et al. 2006). Furthermore, several 16S rRNA gene targeting probes for fluorescence in situ hybridization for the specific detection of different phylogenetic groups of sulfatereducing bacteria are available (Lücker et al. 200). As high throughput method, a DNA microarray targeting all 16S rRNA genes of recognized sulfate reducers was used to characterize the diversity of sulfatereducing communities in pockets and cyanobacterial mats, as well as in peatlands (Loy et al. 2002; Loy et al. 2004). To date, the application of this socalled SRM phylochip was the only study used to investigate specifically the 16S rRNA gene based composition of SRM in wetland systems.

The polyphyletic origin of SRM restricts diversity studies by comparative analysis of their 16S rRNA genes as no single pair of oligonucleotides allows specific detection of all SRM using the

10 Sulfate reducers in peatlands

16S rRNA genes as targets. As alternative, phylogenetic marker targeting functional genes encoding for enzymes involved in the metabolic pathway of sulfate reduction can be used to identify SRM. For instance, the genes encoding for the APS reductase (apsA) as well as for the DSR (dsrAB) were used in several studies to unravel SRM diversity (Hipp et al. 199; Friedrich 2002; Wagner et al. 2005). The presence of both enzymes is not restricted exclusively to SRM, but they share these enzymes with members of the guild of sulfur oxidizing microorganisms (SOM). In contrast to the APS reductase, where apsA of SRM and SOM exhibit a patchy affiliation in comparative phylogenetic analyses (Meyer et al. 200), dsrAB of SRM and SOM are clearly separated in two distinct lineages, allowing a direct assignment of new environmental gene sequences to one of these guilds (Loy et al. 2009). DsrAB genes are present in all known SRM, however, some other DSR carrying microorganisms like Sporotomaculum (Qiu et al. 2003) and Pelotomaculum (Imachi et al. 200), whose dsrAB genes are phylogenetically not clearly separated from those of SRM, are not able to reduce sulfate but live only in syntrophic communities. Beside sulfate, dsrAB carrying microorganisms can use several alternative inorganic and organic sulfur electron acceptors for energy conservation (Rabus et al. 2000). In the absence of a suitable electron acceptor, they are also able to use fermentation to gain energy. Accordingly, dsrAB based surveys identify not exclusively SRM, but include syntrophic microorganisms as well as sulfite, thiosulfate and/or organosulfonates reducers.

Numerous dsrAB based surveys were applied to explore SRM community composition and distribution. For instance, they are well studied in marine ecosystems (Dhillon et al. 2003; Nakagawa et al. 2006; Leloup et al. 200; Leloup et al. 2009), where high sulfate concentrations of up to 2 mM favor their growth and as a result dissimilatory sulfate reduction was shown to be one of the main mineralization processes in marine sediments (Jørgensen 1982). Their widespread occurrence was also demonstrated in brackish systems such as estuarine sediments (Joulian et al. 2001; Jiang et al. 2009), in freshwater habitats such as lake sediments (Karr et al. 2005; Vladar et al. 2008) and in wastewater treatment plants (Schramm et al. 1999; Dar et al. 200). In terrestrial ecosystems, research focuses mainly on specialized waterlogged like rice paddies (Liu et al. 2009), whereas only few studies investigated the SRM community composition in natural freshwater wetlands such as peatlands. For instance, sulfate reducing assemblages in the Everglades were investigated using dsrAB based clone libraries and terminal restriction fragment length polymorphism (TRFLP) analysis targeting the DSR subunit A (dsrA) (Castro et al. 2002; Castro et al. 2005). Sulfate reduction rates revealed higher

11 Part I activities of SRM in eutrophic zones, which were influenced by sulfate input from agricultural runoff, than in more pristine soils (Castro et al. 2002). There, a highly diverse dsrAB composition was detected with the majority of DSR sequences affiliated to different gram- positive Desulfotomaculum spp. and to sequences of yet uncultured microorganisms. In the eutrophic regions, most of the Desulfotomaculum associated sequences were related to microorganisms that are able to completely oxidize their organic substrates, whereas in the pristine regions all Desulfotomaculum-like sequences were exclusively related to incomplete oxidizers (Castro et al. 2002). Additionally, indications were given that sulfate concentration is not the key driver for differences between SRM assemblages, but a selection by the type and amount of electron donor was suggested (Castro et al. 2005). In a more recent approach, spatial relationships between dsrAB diversity and concentrations of metals and sulfur were studied in peatland soils in western New York uncovering a close dependency of dsrAB based community composition and redox stratification. DsrAB genes were mainly detected in deeper soil layers, where sulfur accumulation was evident. In the top peat layers, where manganese accumulation indicated oxic conditions, no dsrAB genes were detected (Martinez et al. 2007).

4. The Schlöppnerbrunnen fen as a model system

The main study site of the first part of this thesis, the minerotrophic fen Schlöppnerbrunnen II, lies in the north-eastern part of Bavaria within the forested Lehstenbach catchment (Fichtelgebirge, Germany). The total sulfur concentration in the fen soil as well as in the surrounding forest and mineral soils is in the upper range of the values reported for forest soils in Europe (~3 – 6.5 g*kg-1) (Prietzel et al. 2009). Most likely these high concentrations are caused from intensive deposition of atmospheric sulfur originating from combustion of soft coal in Eastern Europe. Although there has been a substantial reduction in air sulfur emissions since the late 1980s (Klemm et al. 1999), desorbed sulfate is still transported via groundwater flow from upland soils into the groundwater-fed wetland. Most of the introduced sulfate is retained by accumulation of inorganic and organic reduced sulfur species (RIS and CBS) in the soil (Prietzel et al. 2007), whereby estimates of RIS amounts range from <10% (Paul et al. 2006) to ~35% of total sulfur (Prietzel et al. 2009). Intermediate forms, such as sulfones, are abundant and represent 32% - 41% of the sulfur pool and oxidized sulfur species account for 5% - 23% of the total sulfur pool, respectively (Prietzel et al. 2009). Additionally, sulfate concentrations of soil

12 Sulfate reducers in peatlands solutions demonstrated unexpected stable concentrations over time, which points to a well buffered system regarding sulfate dynamics (Alewell et al. 2006).

Biochemical analysis of several studies led to the conclusion that Schlöppnerbrunnen II is dominated by alternating oxidation–reduction cycles (Alewell et al. 2006; Paul et al. 2006; Prietzel et al. 2009), possibly caused by oxygen diffusion from roots of vascular plants, which are common in the fen (Paul et al. 2006). It has been shown that the habitat is characterized by the coexistence of redox processes, such as nitrate, sulfate, iron, manganese reduction and methanogenesis at a small spatial scale of few cm (Alewell et al. 2006; Paul et al. 2006; Alewell et al. 2008). The presence of high dissolved organic carbon concentrations could favor the simultaneous presence of different reduction processes, as electron donors are possibly not reaction limiting (Alewell et al. 2008). Additionally, in the presence of oxygen, organic and inorganic sulfur species will be reoxidized (Paul et al. 2006) resulting in mobile sulfate formation. Sulfate can to some extent be adsorpted to iron oxide, which is present in high concentrations in the system (Abdelouas et al. 2000; Alewell et al. 2006; Alewell et al. 2008) or is used as electron acceptor by SRM (Loy et al. 2003; Loy et al. 2004; Schmalenberger et al. 200). Part of the sulfate is most likely lost from the fen with the drainage water. Consequently, although it was shown that the Schlöppnerbrunnen II fen can act temporarily as a hot spot for dissimilatory sulfate reduction (Alewell et al. 2001), long-term maintenance of sulfur species within the system is not expected (Paul et al. 2006; Prietzel et al. 2009).

The microbial diversity in the Schlöppnerbrunnen fen systems was examined in several studies. The application of the so-called SRM phylochip was the only study used to investigate specifically the 16S rRNA gene based composition of SRM in the Schlöppnerbrunnen fen system, whereby only Syntrophobacter- and Desulfomonile- like bacteria could be identified. Three further studies investigated the Schlöppnerbrunnen fen system using general 16S rRNA gene targeting oligonucleotides. A two-dimensional single strand conformation polymorphism (SSCP) approach revealed significant differences between the microbial communities of oxic and anoxic zones revealing sequences mainly affiliated to and Acidobacteria as dominant bacterial groups (Schmalenberger et al. 2008). The application of 16S rRNA based stable isotope probing identified, in addition to the aforementioned groups, sequences related to and Clostridia as active anaerobic consumers of monosaccharides like xylose and glucose (Hamberger et al. 2008). Furthermore, uncultured Archaea related to Methanosarcinaceae, Methanobacteriaceae and Crenarchaeota were identified in the heavy

13 Part I fraction of the anoxic slurries. Additional anoxic microcosm studies retrieved further formate consuming and obligate acetoclastic methanogens, related to the families Methanomicrobiaceae and Methanosaetaceae, as potential trophic partners for moderately acid tolerant fermenters (Wüst et al. 2009). Furthermore, dsrAB based clone libraries and TRFLP analyses were engaged to investigate diversity patterns of SRM in the Schlöppnerbrunnen fen. DsrAB based analyses revealed a highly diverse community structure with many novel dsrAB sequences unrelated to known SRM (Loy et al. 2004; Schmalenberger et al. 200). TRFLP analysis distinguished a high diverse community pattern over a depth gradient of 0 cm to 50 cm clustering in three subgroups, namely the upper 10 cm, 15 – 25 cm and 30 – 50 cm depth zones. The environment in the upper fen soil is controlled by the rhizosphere, which permits cation exchange, O2 leakage and release of organic acids. Additionally, due to fluctuations in the water table, the upper layers are exposed to oxygenation and drying, which enhance chemical or microbial renewal of reducible electron acceptors. Thus, detected SRM have to be

O2 tolerant and – as sulfate concentrations were very low – a syntrophic lifestyle of the resident SRM was suggested (Schmalenberger et al. 200).

It was the aim of the first part of this thesis to contribute to a better understanding of the community structure and distribution of dsrAB containing microorganisms, which are possibly involved in the biochemical sulfur cycling in terrestrial wetlands. The description of microbial diversity is the first essential step for the exploration of the composition, interactions, and dynamics of microbial communities within their natural environment. Potent and highthroughput techniques are needed to compile habitat specific microbial inventories and thus, novel tools for rapid community screening, such as a dsrB based DGGE assay and a newly developed functional gene array, were applied.

14 Sulfate reducers in peatlands

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15 Part I

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16 Sulfate reducers in peatlands

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18 Sulfate reducers in peatlands

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19 Part I

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20 Microorganisms with Novel Dissimilatory (Bi)Sulfite Reductase Genes are Widespread in Geographically Distant, Low-Sulfate Peatlands

Doris Steger1, Cecilia Wentrup1#, Christina Braunegger1, Pinsurang Deevong1,2, Manuel Hofer1, Michael Pester1, Michael Wagner1 and Alexander Loy1* 1Department of Microbial Ecology, University of Vienna, Althanstrasse 14, A-1090 Wien, Austria 2Department of Microbiology, Kasetsart University, 50 Phaholyothin Road, Jatujak, Bangkok 10900, Thailand #Current address: Department of Molecular Ecology, Max-Planck Institute for Marine Microbiology, Celsiusstr. 1, D-28359 Bremen, Germany

Draft not corrected by all coauthors

Concept by D.S. and A.L. Microarray analyses performed by D.S. and M.H., DGGE analysis and dsrAB gene library construction carried out by D.S. and C.B. C.W. constructed 16S rRNA gene libraries, P.D. and C.W. performed the qPCR assays. Phylogenetic analyses done by D.S. D.S. and A.L. wrote the manuscript.

Part I

Abstract

Schlöppnerbrunnen peatlands of the Lehstenbach catchment (Germany) house so far unidentified microorganisms with phylogenetically novel variants of the dissimilatory (bi)sulfite reductase genes dsrAB. These genes are generally characteristic for microorganisms that reduce sulfate, sulfite or some organosulfonates for energy conservation, but can also be present in anaerobic syntrophs. The abundance, community dynamics, and biogeography of dsrABcarrying microorganisms in peatlands were unknown. Soils from different depths were sampled at three time points over a six period to analyze the diversity and distribution of dsrABcontaining microorganisms at the Schlöppnerbrunnen II fen site by newly developed functional gene microarray and quantitative PCR assays. Members of novel, uncultivated dsrAB lineages (approximately representing specieslevel groups) (i) dominated a temporally stable but spatially diverse dsrAB community and (ii) represented ‘core’ members (relative abundances of 12%; dsrA copy number versus the total number of bacterial and archaeal 16S rRNA genes) of the autochthonous microbial community in this fen. A bacterial and archaeal 16S rRNA gene inventory of a fen soil sample with relatively high abundance of two novel dsrAB lineages was dominated by clone sequences of the Acidobacteria. Members of acidobacterial subdivisions 1, 2, and 3 were present in the Schlöppnerbrunnen fen at relatively high 16S rRNA gene abundances of 5.626.3%, 615.5%, and 3.613.1%, respectively, with low depthdependent variation, suggesting an important biogeochemical function in oxic and suboxic peat layers. Correlation analysis of quantitative PCR data provided no indications that members of the three acidobacterial subdivisions were carriers of the two novel dsrAB variants. Finally, denaturing gradient gel electrophoresis (DGGE) and clone librarybased comparison of the dsrAB diversity in soils from a wet meadow, three bogs, and five fens of various geographic locations (distance ~1400 km), identified one Syntrophobacter-related and nine uncultivated dsrAB lineages to be widespread in different peatlands. Signatures of biogeography in dsrBDGGE data were not correlated with geographic distance, but could largely be explained by soil pH and wetland type; implying that distribution of dsrABcarrying microorganisms in wetlands on the scale of a few hundred kilometers is not limited by dispersal but determined by contemporary environmental conditions.

22 Sulfate reducers in peatlands

Introduction

Peatlands contain 15% to 30% of global soil carbon (24, 114) and represent a net carbon sink that contributed to global cooling in the past 8,000 to 11,000 years (35). While peatlands are generally rather resilient to external perturbation, longterm global changes such as warming, decreased precipitation, and increased atmospheric deposition of reactive and sulfur compounds might induce transformation of peatlands to new ecosystem types, accompanied by unforeseeable changes in the carbon balance (30). Carbon loss from peatlands is mainly mediated through anaerobic microbial decomposition of organic matter to the greenhouse gases carbon dioxide and methane (55). Ten to twenty percent of globally emitted methane derives from peatlands (15), (119, 121). Primary and secondary fermentation and methanogenesis are considered to be the main carbon degradation processes because of the absence or limited availability of alternative electron acceptors. However, other microbial processes such as denitrification and dissimilatory iron and sulfate reduction can occur together with methanogenesis in the same peat soil fraction and contribute considerably to anaerobic carbon mineralization (63). Fluctuations in environmental conditions on short and longterm scales govern trophic interdependencies among the involved microorganisms. Transitions between synergistic (e.g. syntrophic interspecies transfer of hydrogen/formate) and antagonstic (e.g. competition for the same substrates) microbial interactions determine the extent of carbon flow diversion away from methanogenesis. A prime example is the suppression of microorganisms catalyzing methanogenic carbon degradation by sulfatereducing microorganisms (SRM) that are energeticallyfavored in the competition for substrates such as acetate, alcohols, and hydrogen (36, 115, 116). While sulfate concentrations are generally low in peatlands (10 to 300 M), ongoing sulfate reduction at rates (2.5 to 180 nmol cm3 d1) that are comparable to rates in sulfaterich environments is fuelled by an anoxic recycling of sulfur compounds via the 'thiosulfate shunt' (10). Alternative replenishment of the sulfate pool by reoxidation of reduced sulfur species in the presence of oxygen is dependent on vegetation type and alternating periods with precipitation and drought (31, 92, 124). In addition, increasing global atmospheric sulfur pollution and acid precipitation contributes to terrestrial sulfate pools and is predicted to repress methane emissions from peatlands by up to 15% within the thirst third of this century (37). Given the significance of dissimilatory sulfate reduction, it is surprising that most information about the identity of microorganisms that catalyze this process in peatlands derives from studies of a single model fen system (Schlöppnerbrunnen) located in the forested Lehstenbach

23 Part I catchment (Bavaria, Germany). Different redoxprocesses such as fermentation (43), methanogenesis (44), Fe(III) reduction (100), and sulfate reduction (1, 75) are ongoing on the study site (3). Atmospheric deposition of sulfur originating from combustion of soft coal in Eastern Europe until the 1990s led to the accumulation of sulfur species in the soils of this catchment. Although air pollution decreased in recent years (61), previously stored soil sulfate can desorb and is transported from upland soil into the groundwaterfed fen where it drives dissimilatory sulfate reduction (Alewell 2000). DNA stable isotope probing using in situ concentrations of typical 13Clabeled degradation intermediates (mixture of lactate, acetate, formate, and propionate) has shown that a low abundance Desulfosporosinus species, representing on average only 0.006% of the total bacterial and archaeal 16S rRNA genes, could be responsible for a major part of sulfate reduction in the studied fen (Pester et al. submitted). In addition, other microorganisms that are potentially involved in sulfate reduction were previously detected in this fen using 16S rRNA gene and dsrABbased diversity analyses. Few dsrAB sequences were affiliated with the SRM genera Desulfomonile and Syntrophobacter, while most other sequences could derive from new taxa, as these novel dsrAB lineages have no close cultivated relative (75, 106) Pester et al. submitted). Microorganisms that respire sulfite or sulfate anaerobically depend on the dsrABencoded key enzyme dissimilatory (bi)sulfite reductase for energy conservation and thus these genes have been widely used as markers for PCRbased molecular diversity studies of this guild (29, 58, 67, 95, 117). However, also some organisms that are phylogenetically related to SRM, but have seemingly lost the ability for sulfite/sulfate reduction, can harbor dsrAB. The dsrAB sequences of these organosulfonate reducers (8, 19, 20) or syntrophs (48, 98) can be amplified by the commonly used DSR1F DSR4R PCR primer mix (74). Consistent with the notion that dsrAB can derive from nonSRM, DNA stable isotope probing of dsrAB in the model peatland did not indicate that the novel lineages are linked to lactate, acetate, formate or propionatedependent sulfate reduction (Pester et al. submitted). Besides their unknown ecophysiological function in the fen, additional questions regarding the biology of these enigmatic dsrABcontaining microorganisms remain unanswered: what is their taxonomic affiliation and are they endemic only to this particular fen site or more widely distributed in different types of wetlands? Based on the repeated detection of the same or similar operational taxonomic units (OTUs) of dsrAB (75, 106) we hypothesize that some of these yet unidentified dsrABcontaining microorganisms are part of the autochthonous microbial community in the Schlöppnerbrunnen model peatland. Using newly developed functional gene array (FGA) and quantitative realtime PCR (qPCR) assays, we analyzed the distribution and abundance of individual dsrAB lineages

24 Sulfate reducers in peatlands in different depths of the peatland soil in samples collected at three time points over a period of six years. For linking of dsrAB and phylogenetic sequence information obtained from clone library analyses of bacterial and archaeal 16S rRNA genes, we tested if abundances of selected dsrABOTUs correlated with the abundances of the dominant 16S rRNA gene sequence clusters in the fen. We further investigated the diversity of dsrAB in nine geographically separated wetlands, representing three different wetland types, for patterns in biogeography and endemism by using dsrBdenaturing gradient gel electrophoresis (DGGE) and dsrAB clone library analyses.

Materials and Methods

Description of sites and sampling of soil.

The geographic distribution and major characteristics of the nine wetland sites analyzed in this study are summarized in Table S1 and Figure S1. The main sampling site, the acidic lowland fen Schlöppnerbrunnen II, is located in the Lehstenbach catchment in the Fichtelgebirge mountains (Bavaria, Germany) (see (2, 64, 75) for a detailed site description). Samples collected in 2001 (July 24th), were retrieved from a single soil core, which was divided in four depth sections (07.5, 7.515, 1522.5, and 22.530 cm) (75). In 2004 (September 21st) and in 2007 (May 16th), soil cores (diameter ~8 cm) were collected from three random locations within the site (approximate distance between locations was 15 m) and were subsequently divided in four depth sections (05, 510, 1015, and 1520 cm). Two additional depth sections (2030 and 3040 cm) were collected from two locations in 2007. Besides Schlöppnerbrunnen II, triplicate soil cores (two depth sections ~020 and ~2040 cm) were taken from random locations at eight additional wetland sites in Italy and Austria (Table S1). Immediately at the sampling site, samples for molecular analyses were homogenized, frozen on dry ice for transportation, and stored at 80 °C upon arrival in the laboratory.

DNA extraction.

For microarray, DGGE and qPCR analysis, DNA was extracted from approximately 250 mg (wet weight) of each soil sample using the Power SoilTM DNA Kit (MoBio Laboratories, Solana Beach, CA). For the preparation of a 16S rRNA gene library from Schlöppnerbrunnen II fen soil

25 Part I

(sampled in 2004, 1015 cm soil section), genomic DNA was additionally extracted from triplicate soil samples using the protocol of Zhou et al. (127) and a slightly modified version of the protocol of Griffiths et al. (40). In contrast to the original protocol, precipitation of nucleic acids was performed with 0.1 volume of sodium acetate (pH 5.2) and 0.6 volume of isopropanol for 2 h at room temperature.

Fluorescence labeling of target genes for microarray hybridization.

An approximately 1.9 kb fragment of dsrAB was PCR amplified from 5 ng reference clone DNA or 50 ng environmental DNA using the degenerate primers DSR1Fmix (equimolar mixture of 10 M of each DSR1F variant) and DSR4Rmix (equimolar mixture of 10 M of each DSR4R variant) (Table S2). 16S rRNA and nucleotide transport (ntt) reference genes, targeted by control probes, were amplified by primer pairs listed in Table S3 (50 M of each primer). All forward primers contained a T3 promoter site sequence (5’AATTAACCCTCACTAAAGGG3’) at the 5′ end to enable T3 RNA polymerase-based reverse transcription labeling of the PCR products. PCR mixtures (50 l) were prepared by using 2 mM of each deoxynucleoside triphosphate, 10x

Taq buffer, 2 mM MgCl2 and 2 U of Taq DNA polymerase (Fermentas Inc., Hanover, MD, USA). For amplification of environmental samples, 1 l of bovine serum albumine (20 g/l, New England BioLabs Inc., Beverly, MA, USA) was additionally added to each reaction to enhance PCR efficiency. Reference and control genes were amplified using an initial denaturation step at 95°C for 3 min, followed by 30 cycles of denaturation at 95°C for 30 s, annealing at 48°C (dsrAB), 52°C (16S rRNA gene) or 55°C (ntt) for 30 s, and elongation at 72°C for 1 min 10 sec. The cycling was completed by a final elongation step at 72°C for 3 min. PCR amplification of dsrAB from environmental DNA extracts was carried out by two successive “hot start” PCRs to minimize unspecific amplification products (23). The first PCR was performed in touchdown mode by using an initial denaturation step at 95°C for 3 min, followed by 10 cycles of denaturation at 95°C for 30 s, annealing at 5848°C for 30 s (after each cycle the temperature was reduced by 1°C), and elongation at 72°C for 1 min 10 s. Additional 10 PCR cycles at a constant annealing temperature of 48°C were performed prior to the final elongation step at 72°C for 3 min. PCR products were examined by 1%agarose gel electrophoresis for presence and sizes of amplicons. PCR products were purified from the gel using the Montage™ DNA Gel Extraction kit (Millipore, Bedford, MA, USA) according to the manufacturer’s instructions. One microliter of purified PCR product was reamplified in a second PCR applying an initial denaturation step at 95°C for 3 min, followed by 20 cycles of denaturation at 95°C for 30 s,

26 Sulfate reducers in peatlands annealing at 48°C for 30 s, and elongation at 72°C for 1 min 10 sec. The cycling was completed by a final elongation step at 72°C for 3 min. PCR products were purified using the QIAquick PCR purification kit (QIAgen, Hilden, Germany). Target labeling was achieved by in vitro transcription according to the following protocol. 500 ng of purified PCR product, 4 l 5x T3 RNA polymerase buffer (Fermentas), 2 l dithiothreitol (100 mM), 0.5 l RNAsin (40 U/l) (Promega GmbH, Mannheim, Germany), 1 l each of ATP, CTP, and GTP (each 10 mM), 0.5 l UTP (10 mM), 2 l T3 RNA polymerase (20 U/l) (Fermentas) and 0.75 l Cy3UTP (5 mM) were adjusted with RNasefree water to a total volume of 20 l and incubated at 37°C for 4 hours. The DNA template was subsequently degraded by adding 2 U DNase I (Fermentas) and incubation at 37°C for 15 min. Enzymatic digestion was stopped with 2 l of EDTA (25 mM, Fermentas). Following adjustment of the volume to 100 l with TE buffer (10 mM TrisHCl, pH 7.5; 1 mM EDTA), RNA was precipitated with 10 l of 5M NaCl and 300 l of ethanol. RNA was washed with 500 l of icecold 70% ethanol and resuspended in 50 l TE buffer. RNA concentration and the amount of incorporated dye were measured with a ND 1000 spectrophotometer (Nanodrop, Inc.). Labeled RNA was fragmented by incubating with 10 mM ZnCl2 and 20 mM Tris/HCl (pH 7.4) at 60°C for 30 min. Fragmentation was stopped by the addition of 10 mM EDTA pH 8.0. Labeled RNA was divided in several aliquots that were stored at 20°C.

Probe design and manufacturing of microarrays.

Probes targeting dsrA or dsrB were designed using the probe tools of the ARB software package (78) and a dsrAB ARB database containing approximately 500 sequences (>1500 nucleotides) from pure cultures (74, 129) and environmental studies. Based on a distance matrix tree of all previously published dsrAB clone sequences retrieved from Schlöppnerbrunnen I+II system (75, 106), 146 probes with an average length of 30 bases were designed targeting different clone groups from the study site (Figure S2). Thermodynamic properties of the probes were calculated using the twostate hybridization server of DINAMelt (settings: linear DNA, 37°C, 1 M Na+, 0 M Mg2+, 0.1 mM strand concentration) (80, 81). Probes lengths were adjusted to obtain similar theoretical free energy (deltaG) values. Final probes had a length of 27 to 34 nucleotides with an average deltaG of 41.0 ± 0.7 kcal/mol. The in silico specificity and number of weighted mismatches to nontarget dsrAB sequences of each probe were determined using probeCheck (73). Oligonucleotides for microarray spotting were synthesized by Microsynth (Balgach, Switzerland). The 5’ end of each oligonucleotide probe

27 Part I carried a Tspacer consisting of 30 dTTP molecules and the 5’terminal dTTP was aminated to allow covalent coupling of the oligonucleotides to aldehyde groupcoated VSS25 glass slides (CEL Associates, Houston, Tex.). Each probe was adjusted to a concentration of 50 pmol/l in 50% dimethyl sulfoxide and printed on the slide by using a BioRobotics MicroGrid spotter (Genomics Solutions, Ann Arbor, MI, USA) under constant temperature (20°C) and humidity (min. 50%) conditions. Each microarray contained three and six replicate spots of each dsrA/dsrBtargeted and control probe, respectively. Freshly spotted DNA microarrays were incubated overnight at room temperature in a humid chamber. Slides were further treated with sodium borohydride as described previously (76).

Microarray hybridization.

For clones 80 ng and for environmental samples 500 ng of labeled dsrAB RNA fragments and defined amounts of labeled RNA targeting the control probes (Table S3) were mixed with 120 l of hybridization buffer [0.1% SDS, 0.1% 100x Denhardt’s reagent (Invitrogen), 6x SSC and 15% formamide] and incubated at 65°C for 515 min. The ThermoTWISTER (QUANTIFOIL Instruments GmbH, Jena, Germany) and the HybriWell Sealing System HBW2222FL (Grace BioLabs) were used for hybridization. Microarrays were hybridized for 17h at 55°C under continuous shaking at 400 rpm. Following hybridization, slides were washed by shaking at room temperature for 5 min in 2x SSC/0.1% SDS, for 5 min in 0.1x SSC, and finally for 20 s in ice cold doubledistilled water. Slides were dried by centrifugation (3 min, 300 g), stored at room temperature in the dark, and scanned the same day.

Scanning and data analysis.

Fluorescence images were recorded by scanning the slides at three lines to average and at 10 m resolution with a GenePix Personal 4100A array scanner (Axon Instruments, Molecular Devices Corporation, Sunnyvale, CA, USA). Gain of the photo multiplier tube was adjusted to record images with signal intensities just below the saturation level. Scanned images were saved as multilayer tiff images and analyzed with the GenePix Pro 6.0 software (Axon Instruments). Low quality hybridizations were repeated.

28 Sulfate reducers in peatlands

Microarray hybridization results were first corrected for differences in the local background according to:

−1 SBRPi = S Pi × BPi where SBRPi is the signaltonoise ratio of the probe spot Pi, SPi is the median pixel intensity of the specific probe spot and BPi is the median pixel intensity of the local background area around probe Pi. To account for variations between different hybridizations, the mean signaltonoise ratio x SBRCi of ten internal control oligonucleotides (dsrCONT1 to 4, dsrCONT7 to 12) and differences in labeling efficiency were used to normalize the SBRPi using the following formula:

nSBR = SBR × x SBR −1 ×[dye]−1 ×[template]×100 Pi ( Pi Ci ) where nSBRPi is the normalized signaltonoise ratio of the probe spot Pi, [dye] is the concentration of incorporated Cy3 dye molecules in pmol/l and [template] is the concentration of labeled RNA in ng/l. Heatmaps of the microarray results were generated using the software visualization tool JColorGrid (53). Nonmetric multidimensional scaling ordinations based on BrayCurtis similarities between microarray results (based on the mean nSNRs of all probes in the respective probetarget groups, Figure 1A) were performed using the software PRIMER 5 (http://www.primere.com/) (18).

Quantitative realtime PCR.

QPCR assays were performed using an iCycler IQ Thermocycler (Biorad Laboratories GmbH, München) and the Platinum® SYBR Green I qPCR Super MixUDG (Invitrogen Corporation, Carlsbad, CA, USA) according to the manufacturer’s instructions. 16S rRNA gene and dsrA targeted primers were designed using the “Probe_Design” and “Probe_Match” tools of the ARB program package. Potential formation of primer dimers and theoretical melting temperatures were determined using the open source program Primer3 (http://primer3.sourceforge.net) (103) and the Oligonucleotide Properties Calculator OligoCalc (http://www.basic.northwestern.edu/biotools/oligocalc.html) (56), respectively. Primers, assay performances, and cycling conditions are given in Table 1. Thermal cycling was initiated by a denaturation step at 94°C for 3 min, followed by 4045 cycles of denaturation at 94°C for 40 s, annealing at the specific temperature for 40 s, and elongation at 72°C for 4045 s. PCR products were used as standards for assay calibration and were amplified from the cloned sequences given in Table 1.

29 7 6 5 6 6 6 7 6 a (target (target 10 10 10 10 10 10 10 10 2 2 2 2 2 2 2 2 Dynamic range range Dynamic genes/reaction) 10 10 10 10 10 10 10 10 R² 1.00 Linearity Linearity qPCR performance qPCR (%) Efficiency Efficiency ± 3.083.2 0.98 ± 0.681.5 0.99 ± 2.386.1 0.99 89.8 ± ± 0.489.8 0.99 ± 0.683.1 0.99 73.6 ± ± 1.473.6 0.98 ± 0.177.5 0.99 90.0 ± ± 2.8 90.0 P1K30f P1K30f P5K23f AY167468 AY167480 or clone names) clone or P4K21f, P2K11f P4K21f, X70905, X70906 X70905, X70905, X70906 X70905, AY167478, AY167465 AY167478, AY167467, AY167479, AY167479, AY167467, AY167466, AY167473, Templates for standard standard for Templates synthesis (accession n° n° (accession synthesis P4K3f, P5K18f, P5K16f, P5K18f, P4K3f, P4K23f P4K4f P7K24f et al., al., et submitted Reference Thisstudy Thisstudy Thisstudy Thisstudy Thisstudy Pester Pester 190 330 330 360 160 120 (bp) Approx. length length Approx. of PCR product PCRproduct of 750 250 250 500 [nM] 1000 conc. conc. Primer Primer 69 64 52 68.5 100062.5 360 1000 Thisstudy 100 Thisstudy temp (°C) temp Annealing Annealing and and and related related and OTU6 64 250 OTU2 OTU1 64 64 250 group 1 group 2 group 3 group

bacteria Archaea ntrophobacter ntrophobacter Target group Target Bacteria Bacteria y Acidobacteria Acidobacteria Acidobacteria Acidobacteria spp. S gene dsrA dsrA dsrA Target Target 16S rRNA 16S rRNA 16S rRNA 16S rRNA 16S 16S rRNA 16S content (%) GC(%) length Primer Primer Sequence (5' 3') (5' Sequence imers used for imers quantitativeused realtime PCR. r P

Name Table 1. DsrA243FbDsrA561Rb CACGAC CACCGA TCA GCT DsrA243Fa GTA CTTGGT CAS TTC GGC C DsrA561Ra CACCACCACCGA TGA ACT DsrA216F 18 ATAGGC CTT GGC YGC BAC C 19DsrA561Re CUCGGGAAC 61 AGA CAUCGU U SYBAC836F 18 58 ATAGGC TTT GACCGC TGC 19 C GGGTAC TCA TTC SYBAC986RCTGCTG TG 68 58 19 56 CCGGGG ATG TCA AGC CCA 1389F 19 20 53 1492R 58 55 18 TG TACACA CCG CCC GTTAC GGY CTT GTTACG ACT T 67 16 16 4450 63 Acid303FaAcid303Fb GCGCACGGM CACACT GGA Acid657R GCGCGC GGC CACACT GGAAcid702Fa ATTCCACKC ACCTCT CCC AY Acid702Fb 18 AGATAT CTG CAGCC GAA CAY Acid805R 18 6772 AGATAT CCG CAG GAA CAT CC 19Acid306F 20 78 CTG ATSGTT Acid493RTAGGGC TAG 5060 20 4550 CACGGC CACACT GGC AC AGT TAGCCG CAG CTK CTT CT 50 18 20 18 50 5055 71

30 Sulfate reducers in peatlands

The specific annealing temperature for each primer pair was determined by gradient PCR using perfectlymatching target genes and a selection of nontarget clones with mismatches in the primer target site as templates. For each PCR product, a single band of the expected size was observed by agarose gel electrophoresis. Five nanogram of soil DNA was used as template per PCR. The specificity of each qPCR assay was confirmed by meltingcurve analyses of the samplederived and respective reference clonederived PCR products. Additionally, environmental PCR products were cloned and the identities of a few selected clones were verified by sequencing. Absence of PCRinhibitory substances was confirmed by qPCR analyses (using primers 1389F and 1492R) of dilution series of five selected soil DNA extracts, showing similar efficiencies and correlation coefficients as the standards.

Denaturing gradient gel electrophoresis.

Triplicate soil cores from nine different wetlands (including Schlöppnerbrunnen fen samples taken in 2004) were analyzed using newly developed forward primers for dsrBbased DGGE. DNA extracts obtained from the different depth fractions of one soil core were pooled prior to PCR. Preparation of GCclamp tagged dsrB amplicons was carried out using a nested PCR approach. First, dsrAB fragments were amplified from 20 ng soil DNA in technical duplicates using the degenerate primers DSR1Fmix and DSR4Rmix (Table S2) for hotstart, touchdown PCR as described above, except that only 20 cycles (10 cycles with annealing at 5848°C, followed by 10 cycles at 48°C) were performed. The dsrAB amplicons of duplicate PCRs were mixed, purified by gel electrophoresis using the Montage™ DNA Gel Extraction kit (Millipore), and used as template for the dsrBtargeted PCR. Five different PCR reactions per template were performed using the degenerate primers DSR4Rmix (10 M each variant) and the GC clamp carrying DSR1728Fmixes A to E (Table S5). Reaction mixes (total volume 100 l) were prepared with 2 mM of each deoxynucleoside triphosphate, 10x Taq buffer, 2 mM MgCl2 and 2 U of Taq DNA polymerase (Fermentas), 2 l of bovine serum albumine (20 mg/ml, New England BioLabs) and 2 l of template. Amplification was started by an initial denaturation step at 95°C for 3 min, followed by 20 cycles of denaturation at 95°C for 30 s, annealing at 55°C for 30 s, and elongation at 72°C for 20 sec. Cycling was completed by a final elongation step at 72°C for 3 min. Successfully amplified PCR products of each sample were pooled and concentrated to ~ 50 l using a vacuum centrifuge. Additionally, three dsrAB were subjected to the described procedure and used as an internal standard to allow comparison between DGGE gels. DGGE was performed as described earlier (86). In brief, denaturing

31 Part I gradient of 30 to 80% denaturants (100% denaturants mixture consist of 7 M urea and 40% formamide) was used in an 8% (w/v) polyacrylamide gel. DGGE was performed in 1x Tris acetateEDTA buffer at 60°C and at a voltage of 150 V for 6 h. Following electrophoresis, the gels were incubated for 60 min in a SYBR® Green I solution. For identification, individual bands were excised, reamplified, purified using the QIAquick PCR purification kit (Qiagen), and sequenced directly. Principal component analysis (PCA) (Canoco for Windows 4.5 (112)) was performed using a presence/absence matrix of DGGE bands and environmental variables, i.e. pH, sulfate and nitrate). Schlöppnerbrunnen II was excluded from PCA as corresponding environmental parameters were not determined for these samples. In addition, the DGGE band presence/absence matrix, environmental parameters and geographical data were used for calculation of separate BrayCurtis similarity matrices. The similarity matrices were tested pairwise for similar patterns between wetlands using the RELATE routine of the Primer 5 program (18) (999 permutation, Spearman’s rho).

Clone library construction and comparative sequence analysis.

Bacterial and archaeal 16S rRNA gene clone libraries were constructed from Schlöppnerbrunnen II fen soil (sampled in 2004, 1015 cm soil section) DNA extracts using the primer pairs 616V1492R (77) and Arch21F1492R (27, 76), respectively. Prior to PCR, DNA extracts obtained from the three soil cores by using three different isolation methods (n=9) were mixed in equal parts. Cycling started with an initial denaturation step at 95°C for 5 min, followed by 25 cycles (Bacteria) or 30 cycles (Archaea) of denaturation at 95°C for 40 s, annealing at 52°C (Bacteria) or 56°C (Archaea) for 40 s, and elongation at 72°C for 1 min 30 sec. PCR was completed by a final elongation step at 72°C for 10 min. After TOPO TA cloning (Invitrogen) and sequencing, sequences were checked for chimeras with the software tools Bellerophon (46) and Pintail (http://www.bioinformaticstoolkit.org/WebPintail/) (5) and by comparing the phylogenetic placement of different parts of each 16S rRNA gene sequence (i.e. base positions 26747 and 7471471, numbering according to E. coli). A dsrAB clone library was construction from pooled DNA extracts of all Rasner Möser fen soil subsamples using the degenerate primers DSR1Fmix and DSR4Rmix (Table S2). Touchdown PCR was performed as described above, with the exception that 25 cycles (10 cycles with annealing at 5848°C, followed by 15 cycles at 48°C). The cycling was completed by a final elongation step at 72°C for 10 min. The presence and sizes of the amplification products were

32 Sulfate reducers in peatlands determined by agarose (1%) gel electrophoresis. Prior to cloning using the TOPO XL cloning kit (Invitrogen), PCR products were purified from the gel using the Montage™ DNA Gel Extraction kit (Millipore). Operational taxonomic units definitions and Chao1 (17) estimates of richness were carried out using the software package DOTUR (105). Coverage was calculated according to the method described by Good (39). Phylogenetic analyses were performed by using distance matrix, maximumparsimony and maximumlikelihood methods implemented in the ARB program package (78). 16S rRNA sequences were analyzed using the SILVA 16S rRNA database SSU Ref version 98 (97) and the SILVA Web Aligner SINA was applied to identify the two next related sequences of each fen soil clone. The bacterial 16S rRNA tree was inferred using PHYML and a 50%conservation filter for Bacteria (1,248 valid alignment positions). The iTOL tool was used for tree visualization (69). The archaeal 16S rRNA tree was calculated using TreePuzzle (111) and a 50%conservation filter for Archaea (1,229 valid alignment positions). Deduced DsrAB and DsrB amino acid sequences were grouped in OTUs based on an identity threshold of 90% (approximately defining specieslevel groups, (58, 75). Phylogenetic analyses were performed using an indel filter (543 valid alignment positions after exclusion of variable regions with insertions and deletions). A consensus tree was drawn based on PHYLIP ProML (JTT), protein parsimony, and distancematrix (FITCH, JTT, global rearrangements, and randomized input order of sequences) analyses of sequences >500 amino acids. Sequences <500 amino acids were individually added to the trees without changing the overall topology by using of the ARB "parsimony interactive" method.

Sequence accession numbers. The sequences determined in this study have been deposited in the GenBank database under accession numbers GU127834 to GU127875 (archaeal 16S rRNA gene clones), GU127668 to GU127833 (bacterial 16S rRNA gene clones), GU127936 to GU127971 (dsrAB clones), GU127876 to GU127935 (dsrB clones).

33 Part I

Results

Temporal and spatial dynamics of dsrABcontaining microorganisms in the Schlöppnerbrunnen II fen. A habitatspecific FGA, consisting of 146 oligonucleotide probes, was developed and optimized for diversity analysis of dsrABcontaining microorganisms in the Schlöppnerbrunnen fen system (Table S4, Figure S2) (75, 106). In a first step, specific hybridization conditions were established by performing a series of hybridizations with clone dsrSBI3 (AY167467) at increasing formamide concentrations (0 to 60%, at 5% increments) in the hybridization buffer. A formamide concentration of 15% was selected for all subsequent hybridizations as the best compromise between sensitivity (i.e. high signal intensity of perfectly matched probes) and specificity (i.e. high signal intensity ratios between perfectly matched and mismatched probes). In the second step, the specificity of the dsrABFGA was further evaluated by individual hybridization with 36 dsrAB reference clones. This set of reference clones contained at least one perfectlymatched target for 120 probes. 26 probes could not be fully tested, because perfectlymatching reference clones were not available. 98 of the 120 tested probes gave a positive signal only with the perfectlymatching target clone. False positive hybridizations only occurred between probes and nontarget sequences with less than two weighted mismatches. Two probes (dsrB40 and dsrA127) were false negative and were thus excluded from further analysis. In summary, only 0.57% and 0.04% of the 5256 individual probetarget hybridizations were false positive and false negative, respectively (Table S6). The nSNR values of perfectly matched probetarget duplexes of the final probe set ranged from 0.04 to 2.2 (factor 55), showing that the signal intensities of individual probes varied considerably. In the third step, the sensitivity of the dsrAB FGA was assessed by a series of hybridizations containing different concentrations of the clone dsrSBI36 (AY167469). One copy, 10, 100, 1000, 10000, and 100000 copies of the were added to 60 ng (corresponding to approximately 1x107 microbial ; assuming an average size of 5 Mbp) aliquots of fen soil DNA (sampled in 2007, depth 1520 cm) prior to PCR amplification and further target preparation. A minimum of 1000 plasmids was required for positive detection of clone dsrSBI36 by dsrABFGA hybridization. Given that most dsrABcontaining microorganisms only have a single copy of dsrAB in their genome, this corresponds to a detection limit of 0.01% of the total microbial community for this target

34 Sulfate reducers in peatlands organism (nSBR values of probes targeting clone dsrSBI36 ranged from 0.170.41 in the specificity ). Additionally, the spottospot variability (from triplicate spots of the same slide) and the slidetoslide variability (from replicate spots of different slides) were consulted to evaluate the robustness of the microarray analysis. The spottospot variability of five randomly chosen arrays, given as mean coefficient of variation of SBR values, was 3.8 ± 0.7% between triplicate spots on a given slide and 7.4 ± 3.7% between spots on three separate slides hybridized with the same fluorescently labeled target RNA. Technical variability of environmental microarray analyses, as determined by hybridizations with labeled RNA prepared from the same environmental sample (year 2001, depth 010 cm) but from separate DNA extractions and labeling reactions, showed a mean coefficient of variation of 15.3% based on nSBR values. This variability includes the composite methodological biases introduced by DNA extraction, PCR, labeling, and hybridization.

The newly developed Schlöppnerbrunnen fenspecific dsrABFGA was applied to analyze the spatial and temporal distribution of dsrAB at the site Schlöppnerbrunnen II. Microarray analyses of soil samples obtained from different depths (0 to 40 cm) were performed in technical triplicates (i.e. analysis of three separate DNA extracts obtained from the same soil core) for the year 2001 and in biological triplicates (i. e. analysis of three soil cores) for the years 2004 and 2007. Microarray procedures for target preparation, hybridization, and data analysis were kept identical for all soil samples to allow comparison between different hybridizations. Soil depth profiles of dsrABFGA hybridization patterns that were obtained for the three years were generally similar (Figure 1A) average BrayCurtis similarity of 59±18% among all samples). However, in deeper soil layers the dsrAB diversity was considerably greater than in the top soil layer, where only a few probetarget groups were detected by the FGA. In accordance with this observation, multidimensional scaling (MDS) analysis of BrayCurtis similarities between microarray hybridization patterns clearly separated the top soil layers from all other samples (Figure 1B). Members of the novel dsrABOTUs 1 and 4 (75), which are only distantly related to Desulfobacca acetoxidans (DsrAB identities of 6977%), were present in most deeper soil fractions, as indicated by positive probe signals for probetarget groups 1 and 29, respectively.

35 Part I

A 8 7 9 1 2 3 4 5 6 11 14 15 17 18 20 21 24 28 29 31 34 35 40 43 46 49 52 55 56 57 10 13 16 19 22 23 25 26 27 30 32 33 36 37 38 39 41 42 44 45 47 48 50 51 53 54 58 2001 12 0 – 10 cm 10 – 17.5 cm > 0.39 17.5 – 25 cm 0.36 0.39 25 – 30 cm 0.33 0.36 0.30 0.33 2004 0.27 0.30 0 – 5 cm 0.24 0.27 5 – 10 cm 0.21 0.24 10 – 15 cm 0.18 0.21 15 – 20 cm 0.12 0.18 0.12 0.15 2007 0.09 0.12 0 – 5 cm 0.06 0.09 5 – 10 cm 0.03 0.06 10 – 15 cm < 0.03 15 – 20 cm 20 – 30 cm 30 – 40 cm OTU6 OTU3 OTU7 OTU2 OTU1 OTU8 OTU9 OTU5 OTU4 OTU11

B

Stress: 0.04 2030 cm 3040 cm>83% 1520 cm >55%

1015 cm 2530 cm 1017.5 cm

17.525 cm

510 cm 510 cm >51% 1015 cm >71% 1520 cm 05 cm 010 cm

05 cm

20012004 2007

Figure 1. Microarraybased dsrAB diversity analysis of Schlöppnerbrunnen II fen soils sampled from different depths in the years 2001, 2004, and 2007. (A) Results of microarray hybridizations are displayed as mean nSBR of all probes within a probetarget group and are averaged between triplicate hybridizations (results of individual replicate analyses are presented in Figure S3). The color code translates into different mean nSBR values. Probetarget groups are arranged phylogenetically according to their position in schematic dsrAB neighborjoining tree. Affiliation of probetarget groups to previously determined dsrABOTUs (75) is indicated. (B) Multidimensional scaling plot based on BrayCurtis similarities between microarray hybridization patterns shown in panel A. Minimal BrayCurtis similarities of all samples in a given group of samples is indicated in color.

36 Sulfate reducers in peatlands

The following dsrAB groups with no close, cultivated relatives were also detected frequently in some of the deeper soil layers: Members of OTU 2 (probetarget group 49) and the related probetarget groups 45, 46, and 47, probetarget groups 41, 42, and 43, members of OTU 3 (probetarget groups 3133) and OTU 11 (probetarget groups 1315). Microorganisms related to the genera Desulfomonile (probetarget group 28) and Syntrophobacter (OTU 6; probetarget groups 21, 23, 25) were only occasionally detected by dsrABFGA analysis.

For confirmation of microarray results, dsrAtargeted qPCR assays for OTU 1 (probetarget group 1), OTU 2 (probetarget group 49), and OTU 6 (probetarget groups 21, 23 and 25) were developed and applied to determine the abundance of these OTUs in the Schlöppnerbrunnen II soil samples. The average total number of bacterial and archaeal 16S rRNA genes for all samples from different soil depths and years was 3.6±1.6 x 107 copies per gram of wet soil, with total numbers being slightly higher in upper soil layers (Figure 2).

Copy number ratios Relative abundance (%) OTU1 OTU2 OTU1 OTU6 OTU6 OTU2 OTU 1 OTU2 OTU 6 16.7 2.0 8.6 2001 0 – 10 cm 0.05 0.01 0.00 116.4 25.4 4.6 10 – 17.5 cm 1.05 0.23 0.01 87.0 93.8 0.9 17.5 – 25 cm 1.50 1.61 0.02 25 – 30 cm 8.0 5.7 1.4 0.41 0.29 0.05

30.5 1.9 16.3 2004 0 – 5 cm 0.09 0.01 0.00 6.9 2.5 2.7 5 – 10 cm 0.09 0.03 0.01 46.6 34.6 1.3 10 – 15 cm 1.27 0.94 0.03 5.8 14.0 0.4 15 – 20 cm 0.48 1.14 0.08 18.5 12.4 1.5 2007 0 – 5 cm 0.05 0.04 0.00 16.1 31.7 0.5 5 – 10 cm 0.09 0.17 0.01 8.0 42.0 0.2 10 – 15 cm 0.26 1.35 0.03 2.2 48.1 0.0 15 – 20 cm 0.08 1.70 0.04 2.5 26.6 0.1 20 – 30 cm 0.11 1.22 0.05 30 – 40 cm 1.8 17.4 0.1 0.07 0.69 0.04

1 10 100 1000 104 105 106 107 108 109 Gene copy number per gram of wet soil

Figure 2. Absolute and relative abundances of three dsrAB-OTUs in different depths of the Schlöppnerbrunnen II fen site in the years 2001, 2004, and 2007, as determined by qPCR. Error bars are the standard deviations of the mean for the three replicates. Black circles indicate total number of bacterial and archaeal 16S rRNA genes. Copy numbers of dsrA of OTUs 1, 2, and 6 are displayed as white, grey, and dark grey bars, respectively. Additionally, the relative abundance of each dsrABOTU (given in % of the total number of bacterial and archaeal 16S rRNA genes) and copy number ratios between individual dsrABOTUs are shown for all years and depths.

37 Part I

In contrast, dsrA copy numbers of all three target OTUs were lowest in the top soil layers and generally increased with soil depth, although OTU 1 and OTU 2 were represented by slightly lower dsrA copy numbers in the deepest soil layers. In all years and soil depths, dsrA of the novel OTUs 1 and 2 outnumbered dsrA of the Syntrophobacterrelated OTU 6; with ratios ranging from 1.8 to 116.4 for OTU 1 and from 1.9 to 93.8 for OTU 2 (Figure 2). The mean dsrA copy number per g fen soil was 8.9 ± 0.6 x 104, for OTU 1, 2.1 ± 0.07 x 105 for OTU 2, and 7.9 ± 0.6 x 103, for the OTU 6. The mean relative abundances (percent ratio of dsrA copy numbers to total bacterial and archaeal 16S rRNA gene copy numbers averaged over soil depths) of OTUs 1, 2, and 6 were 0.75%, 0.53%, and 0.02% in the year 2001, 0.48%, 0.53%, and 0.03% in the year 2004, and 0.11%, 0.86%, and 0.03% in 2007, respectively. For Syntrophobacterrelated bacteria (represented by OTU 6), an additional 16S rRNA gene targeted qPCR assay was developed based on previously published 16S rRNA gene clone sequences from the Schlöppnerbrunnen fen sites (75). As expected, 16S rRNA gene and dsrA based qPCR data of the Syntrophobacterrelated target group showed a high correlation (Pearson correlation coefficient of 0.854, p<0.01), confirming that dsrAB of OTU 6 indeed derived from Syntrophobacterrelated bacteria in the Schlöppnerbrunnen fen soil samples.

Comparisons of dsrA copy numbers to the nSBR of the corresponding probetarget groups of the dsrABFGA demonstrated significant correlation (p<0.01) between qPCR and microarray data; with Pearson coefficients of 0.811 for OTU 1 (with probetarget group 1), 0.756 for OTU 2 (with probetarget group 49), and 0.734 (with probetarget group 21) and 0.863 (with probe target group 23) for OTU 6.

Bacterial and archaeal 16S rRNA gene inventories of the Schlöppnerbrunnen II fen. Soil sampled in the year 2004 from 10 to 15 cm depth contained members of the novel OTUs 1 and 2 at a relatively high abundance and thus was chosen as a representative sample for phylogenetic identification of the abundant Schlöppnerbrunnen II fen soil . A bacterial and an archaeal 16S rRNA gene clone library were constructed and phylogenetically analyzed. After removing chimeras from the bacterial (n=9) and the archaeal (n=1) dataset, 16S rRNA gene sequences with identities ≥99% (approximately defining species level phylotypes; (109)) were grouped in OTUs/phylotypes.

38 Sulfate reducers in peatlands

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clone P8K on BacB 007 ne F K 89 9 c S 038 co 5K (1) s B 7 b pe C12 56 2 5193 2 (2) f P4K 2 3 l Ca SP 5 u e ores K 23 i P oil clo o P0 9 soil 1 AJ231195 Isosphaera pallid str. e B 4 lone 0f 132 J58 38 il c f o obic 8 f F3 7 fen soil clone P7K18f 33 e DQ6 3 C AY622271 subsur s mir fen s P7 Q W4 A Q1 cin 1 f fen soil clone P2K10f 2 s er A il c K t lo 1 fen Q1 47 u 2 D n lon a 7 (1) group 3 V so K 1 EU 76 s (1) AKIW863 ne gar ne P iphila 5 D c t eSm fe 2 so P oil cl 17 f 40 oil contam 1f (1) il X71838 3 2 cl FC 7 fen soil clone P8K21f be olyticu 951 oil o DQ984569 soilfen clone soil clone IFD 124 P9K2 f DQ138961 ricen straw soil clonepaddy soils clone JG29539 one o 7 DQ A o n PT43 K 1 fe 4810 778 E AJ390478 rice roots clone PR 3 n 6 n e 1 DQ088e Sar U135 e F 335 il cl X62911 Planctom F f f o (2) P s E U 28 (1) A 17f s 286 g fen soil clone P1K3 6K C 2 E 7 (1) EU266 group 15 4 62 il clone P1K 13 fen o E ra 7 2 120 F0 ss pra f CP000473 Solibacter usitatus Ellin6076 W 3276 16S 1224 AB (1) fen s e F 7 fen soil a ment clone 35 19 n 1A 8 cy AY963300 forestirie s soil clone A fen soil clone P4K21f lo 1 31 yces lim 9f AY326547 Amazon soil clone 55 (1) c e 36 ano nd on G (1) oil clo EU335400 soil clone BacC u 028 cl S J bacte clone P2K7f DQ129640 urban aerosol clone etla il 13 fe nophilus fen soil clone P7K19f w so 11 F/G EF rial m n ne 1f P7K23f e C 494 so FFC R fen soil clone P2K11f ed c E e K il 7 st fa e lon 37 at c c DQ833485 sedi re ur f (1) lon c 5 2 lo H284 fo s 24 ile Riv lone ne AF465658 thermal8 soilu bcloneK YNPRH5il c p B 1 er P 8 s 5 so ste 045 P gra HAV 1 EU33537339 soil clone2 BacBP re096 (1) a su 050 n (1) K2 1 fen soil2 clone87 ne tu 23f g w Ns SM DQ303 itic f F5 6 lo as 5K U H la fen s Omat6H04 A 90 il c p P inin e D ne 336 Tu U nd Q o 03 ne m n o 35000 sc oil c D n s 3 m lo A ap 8 fe 83 niu t soil cl Y9 ber m B e lone 0 ra t soil c 13 rady clo P9 Q s (2) e pea 524 ag rhiz ne D n soil clo57 u re 0 fo natum clone N K2 fe 5 fo u mir 18 res ob R 3 2956 26 ssu t so 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clone B76 13376 forest soil clo sphagnum pea ne DUNssu169 AM162428 acidic (2) fen soil clone P4K25f fen soil clone P5K27f (1) AY234551 soil isolate Ellin5134 DQ451505 forest soil clone FAC66 lev 16S 1212 AB262724 Sarobetsu m F019854 aspen rhizosphere clone E ire peat soil clone HSM050P B 13 E brifaciens fen soil clone P4K23f D86512 Acidisphaera ru (1) D2 6171 Acidobacte e RB215 fen soil clone P9K25frium capsulatum DQ386264 Siderooxidanssp lithoautotrophicushere clon ragmites rhizo e P2K2f EF5158 (8) 0323 Ph fen soil clon fen soil 77clone gras P5K16fslan AB24 ortensis 10% d soil clo (5) 11f ne FCPT602 omonas he P3K EU150198 spr echlor il clon 6 (10 AY277621 D fen so SS 9 AY96 uce fir fo clone fe 3461 rest clo (4) ctor 5K33f n soi forest soil c ne NiA biorea ne P fe l clone lo SF 9 eded il clo 6059 n so P5K ne BS e se fen so e 65 fen s il clo 18f 23 sludg lon oil c ne P5K3 (1) 5900 (1) ent c lo 1f AY94 edim 2K1f EU3 ne P (2) ted s ne P fen s353 3K1f ina I 27 oil clo67 s (6) tam il clo X AB238777 oil c con HAB ne P4Klon 4614 fen so AB262723 Sarobetsu miree Bac peat soil clone HSM050P B 1 DQ40 (1) ne C 13f 3f B 0 e clo P2K fe Sarobetsu (6)Mire peat55 soil clone HSM SS 0 rin ne n soil 8 ma cidia frumentensislo fen c 4091 il c WD260nes soil clolon AF526927 Mars OdysseyJ2 encapsulation facilityn so clone T5 3ge 17 EU335320 so e P4 A fe li 2 K lca 4 5f AY ne P1 2 (2) d soil cloneas a D1 3K fen 39 f on ne P 1 5 K30f(3) clo ne 4b s 450 p il clone BacB 043 dom oil EU oil c eu s il clo e K fen 33 lon ast (2) dy so clon 65596317f s 531 e P ure 14 AJ292673201 pollute Ps pad fen K AY oil c 9 so s CQ778955 Candidatus Endoecteinas40 ne P4 f 963 3K oi U2 ded t bog olinii72 en lone il clo 2f l c E o (1) f E s 394 lone 2 flo pea ent clolone A 1 0 U o P5 (1) 1 il c J 2 AY9633416 il forest fosoil clone AH48ne E 783 num im o S K 8 cl K1 BacB 0 B113 g ed s e 2 fen soil clone0 P3K14fon re J61 n P 46 e st soil 1f 1 A ed s fen ne f EU150219 spruce1 fir forestP clon 7 spha at clo fen soil clone P1K14ffo 4 (1) 35 5 (1) res K1f clo rtium fen soil clone P3K4 t s ne F5248 ntamin fen soil clone P4K28f (4) AS A nso soil clo oil 4 co o fen AY963454 clo 26 83 X70905g c Syntrophobacter w fen soil clone P5K3f n in DQ451448 forest soil clone FA e S rm (2) 9 DQ404 fo EU680451 forest soil clone S5 169 1 DQ451447 forest soil clone (1) 3 46 ans fen soil clone P5K6 7f fe forest soil clone BS16 1 7 n soil c tr y clone Er LLAYS 99 F E B D F C (1) ithella propionica 1K 2 f (7) T L42613 Desulforhopalus vacuolatus f fen Q 51 2 e EU clone mbI eb P 489 4 6 (1) 20 A 5 e NiA SF 31 ent clone 655973n on S fe soil clo 2 lo Acidobacteria 950 AF200962 arctic isolate LSv22 l B2 150 071 03 (1) m c tepidum 55 EU234214 river clone C3 n group1 fen soil clone P1K24f 5K n J00 il CP acilis A e P A r U 1 o P 04 fen soil clo Y so 5 gr (1) AC 20 0 2 AJ831750 chemocline clone 172 oil clos g 4f X92709 peat bog75 clone21 TM232 8 1 tu as 3 (3) e F r F F f D il c ne AB K8 AF126282 Sm d n 8 2 3 l te fe Q 647 ne n sl n s S D 41 5 L spru EU542537 soil slurr K8 f fe lo dra 9K2 is A a f fe fe n 45 ne AS 9 D Q 26 P5K22 fie E fen soil clone P7K25f n 6 A 2 1 AB AX814126 D inated sedi aliangium lone 1B P J n s 91 be n il clo bac 17 a (1) m f n s A d s Q129128 forest soil clone 6 4 c lo 0 8 U 53 e P me 6 P5 M943 1513 fore dy oil clo o c 02 71 s oil 2 5 c d h 6 tob C 11 c e 2 D o sto ne 14f 0 72 e (1) oil dens 5K S 07 o 9 y ne oil clon 6 id 6 fir fo FAC 9 il 67 a i pa s P V U clo l 87 e B nze nd s on lo 7 2 ne P9K22fim K a d l c f c fen s clo MO 7K e C 24 tu ow s n c e M 16 S 1 l la t soil c c e RB 6 l c 0 ana 5 o HS 6 o ne P illu (1) 8 est l ne i 1f s n ri il c n 4 c n 1 9 AB426191 lotus field soil ss e n aro ne c rest clo o 9 r e 1 oi o DK 7 u soil 0 4 l lo n e s clo o for b o AB062751 H (1) 8 ne r st soil clo F fen soil clone P6K12f c e P5K n ore s s c b (1) v EU134562 prairie soil clone FFCH3849 n A c Ca a il 31 6 4 l P 1 b C n or dra il clo iu cter c i og niu ontam itu 7 e a c 3 gra 3 f fe ehalococcoides ethenogen 7 K e PP 90 re a t clo o m lo 89 fe n rb 49 ldilin e t DQ404843 contam 7 2 K2 13f lin 5 so s s n o t b m ne 04 1 DQ (1) 3 l al fo m m f 4 he on u e NiA U en c xib n oil clo u 03 10f (1) er p 7f m F0 E 16 45 633 ea s e s 86 (1) le fe s ina a 5 ted (2) e r J adow s (2) p d n o ke r i F dim c inin a (2) re 2 DQ A e e FA o SF E e ted

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Figure 3. 16S rRNA genebased maximum likelihood (PhyML) tree showing bacterial sequences retrieved from Schlöppnerbrunnen II fen soil. The tree includes for each OTU a representative clone (shown in bold), sequences from the next closest relatives, and representative cultivated microorganisms in the ARBSILVA database. For each OTU the number of retrieved clones is depicted in brackets. For the phylum Acidobacteria, the grouping of subdivisions 1, 2, 3, and 15 (nomenclature according (7)) is shown. Scale bar indicates 10% estimated sequence divergence.

39 Part I

The 166 bacterial and 42 archaeal clones formed 85 and 18 OTUs and covered 69% and 76% of the expected diversity, respectively. Chao1 richness analyses estimated a minimal number of 160 bacterial and 25 archaeal OTUs. The bacterial OTUs affiliated with members of the eleven described phyla Actinobacteria, Acidobacteria, Bacteroidetes, Chloroflexi, Chlorobi, Elusimicrobia, Firmicutes, Nitrospirae, Planctomycetes, Proteobacteria (including alpha, beta, gamma and deltaclasses), and Verrucomicrobia, and the uncultured candidate phylum TM6 (Figure 3). Most sequences (53.6%), corresponding to 33 OTUs, derived from the phylum Acidobacteria. Most OTUs (n=77) showed low similarity (<90%) to sequences of cultured members of the respective phyla. Only 8 OTUs, represented by the following clones, had 16S rRNA sequence identities of >95% (approximately defining genus level phylotypes): P2K20F (99% to Desulforhopalus vacuolatus, L42613, Deltaproteobacteria), P9K23f (97.5% to Bradyrhizobium elkanii, U35000, Alphaproteobacteria), P5K6f (96.1% to Smithella propionica, AF126282, Deltaproteobacteria), P2K2f (95.78% to Siderooxidans lithoautotrophicus, DQ386264, ), P6K11f (95.4% to Desulfosporosinus orientis, Y11570, Firmicutes), P1K21f (95.2% to Solibacter usitatus Ellin6076, AY673303, Acidobacteria), P4K25f (95.1% to isolate Ellin5134, AY234551, Alphaproteobacteria), and P3K5f (95% to Syntrophobacter wolinii, X90605/06, Deltaproteobacteria). The next relatives to the recovered 16S rRNA sequences of this study originated from a broad range of habitats. However, the majority of sequences (68%, n=87, corresponding to 64 OTUs) were closest related to sequences from soil, with several clones (n=29, corresponding to 20 OTUs) being related to sequences obtained from wetlands or watersaturated soil habitats e.g. forested wetland impacted by reject coal (14), rice paddy field (89), peat bogs (26, 102), alpine tundra and wet meadows (21). Furthermore, 46 sequences (28%) had 16S rRNA sequence identities of >95% to sequences retrieved from heavy DNA fractions after stable isotope probing of Schlöppnerbrunnen II fen soil mesocosms that were incubated with and without sulfate (ZITAT MICHA). Most sequences were in common to sequences obtained from both incubations with and without sulfate and affiliated to Acidobacteria (n=36, 95.1% to 99.7%), Planctomycetes (n=1, 96.4%), Actinobacteria (n=1, 95.1 to 95.5%) and Alphaproteobacteria (95.4%). However, similar Nitrospirae sequences (n=6, 97 to 99.4%) and Firmicutes sequences (n=1, 96.6 to 97.6) were detected only in incubations with sulfate. With the exception of acidobacterial clones P5K31f (98.2% to AM270582) and P1K30f (99% to AM773947) and the Chlorobi clone P5K17f (96% to AM773938), most sequences from our study represent genus level phylotypes that have not been detected by three other 16S rRNA based studies of the Schlöppnerbrunnen fen system (43, 75, 125).

40 Sulfate reducers in peatlands

72 DQ278152, soil clone RotF135iia 80 DQ278155, soil clone OdenF150iiia 100 DQ278153, soil clone OdenF150iia Schlöppnerbrunnen clone A_P1K6f (2) 76 100 77 Schlöppnerbrunnen clone A_P2K26f (1) Schlöppnerbrunnen clone A_P2K16f (3) 100 Schlöppnerbrunnen clone A_P2K32f (10) 99 97 AB243803, rice paddy field clone NRPL AB243793, rice paddy field clone NRPB Schlöppnerbrunnen clone A_P1K8f (2) 88 100 100 DQ901270, acid copper mine clone K1G43 AB262710, Sarobetsu mire clone HSM050PA9 DQ223194, subsurface water clone EV221H2111601H196 95 90 DQ223192, subsurface water clone EV221H2111601H168 94 100 Schlöppnerbrunnen clone A_P2K3f (2) Schlöppnerbrunnen clone A_P2K18f (1) AB262708, Sarobetsu mire clone HSM050PA7 AB243795, rice paddy field clone NRPD Schlöppnerbrunnen clone A_P2K15f (1) 91 EU239960, Candidatus Nitrosocaldus yellowstonii EU281336, Candidatus Nitrososphaera gargensis 85 AB294263, groundwater clone SWA07 Schlöppnerbrunnen clone A_P2K11f (9) AB050208, gold mine clone SAGMAD EU309868, rhizosphere clone LAR_Cren_60 AJ576209, leachate landfill clone GZK8 97 EU280219, salt marsh sediment clone HQSAT_10F8 96 Schlöppnerbrunnen clone A_P2K21f (1) 98 Schlöppnerbrunnen clone A_P2K14f (2) Schlöppnerbrunnen clone A_P2K30f (1) Schlöppnerbrunnen clone A_P3K4f (1) Schlöppnerbrunnen clone A_P2K20f (1) U59986, sediment clone pGrfC26 AJ548934, oligotrophic fen cloneFenO16S EU155995, rich minerotrophic fen clone MH1492_3B 76 EU155994, rich minerotrophic fen clone MH1492_12C Schlöppnerbrunnen clone A_P3K3f (1) EU155991, rich minerotrophic fen clone MH1492_4A EU155993, rich minerotrophic fen clone MH1492_B9G 100 DQ060322, Ignisphaera sp. Tok10A.S1 Crenarchaeota X14835, Thermofilum pendens 100 EU155939, rich minerotrophic fen clone MHLsu47_B8A 100 Schlöppnerbrunnen clone A_P2K23f (1) 100 EU155938, rich minerotrophic fen clone MHLsu47_B11B 100 CP001338, Candidatus Methanosphaerula palustris E19c CP000254, Methanospirillum hungatei JF1 98 100 EU155965, rich minerotrophic fen clone MH1492_B8C 100 Schlöppnerbrunnen clone A_P1K22f (1) 99 EU155964, rich minerotrophic fen clone MH1492_B6H 100 X16932, Methanosaeta concilii 100 Schlöppnerbrunnen clone A_P2K10f (2) EU155955, rich minerotrophic fen clone MH1492_B9E EU155912, rich minerotrophic fen clone MHLsu47_22D 10% Euryarchaeota

Figure 4. 16S rRNA genebased maximum likelihood (TreePuzzle) tree showing archaeal sequences retrieved from Schlöppnerbrunnen II fen soil. The tree includes for each OTU a representative clone (shown in bold), sequences from the next closest relatives, and representative cultivated microorganisms in the ARBSILVA database. For each OTU the number of retrieved clones is depicted in brackets. TreePuzzle support values over 70 are given at the respective nodes. Scale bar indicates 10% estimated sequence divergence.

Most archaeal OTUs (n=15) were affiliated with the phylum Crenarchaeota, while only 3 OTUs branched within the Euryarchaeota (Figure 4). While none of the crenarchaeal OTUs from the Schlöppnerbrunnen II fen was closely related to a cultivated microorganism (16S rRNA sequence identities <80%), most OTUs grouped with other soil and wetlandderived sequences. Clone A_P2K11f, representing an OTU consisting of nine sequences, was affiliated

41 Part I with Nitrososphaera gargensis, an archaeum recently proposed to belong to a novel archaeal phylum, the Thaumarchaeota (13) (Spang et al. submitted). The euryarchaeal OTUs were closely related (sequence identities of 9899%) to clones from a rich minerotrophic, pH neutral fen located in central New York State (16). The closest described relatives of clones A_P2K23f and A_P2K10f were Candidatus Methanosphaerula palustris and Methanosaeta concilii (each with a sequence similarity of 94%), respectively. In contrast to the bacterial sequences, a considerable proportion of archaeal sequences (43%) had highest similarities to sequences from a previously published 16S rRNA gene inventory of the Schlöppnerbrunnen fen system (125).

Abundance of Acidobacteria in the Schlöppnerbrunnen II fen. Sequences of the phylum Acidobacteria predominated in the bacterial 16S rRNA gene library. Development and application of specific 16S rRNA genetargeted qPCR assays for the three main acidobacterial subdivisions/groups 1, 2, and 3 (Fig. 3, Table 1) confirmed that each of the three groups constituted a major part of the Schlöppnerbrunnen II fen soil microbial community in all soil depths and years (Figure 5).

Total bacterial and archaeal 16S rRNA gene copy number per gram of wet soil 1 10 100 1000 104 105 106 107 108 2004 0 5 cm 5 10 cm 10 15 cm

15 20 cm 2007 0 5 cm

5 10 cm 10 15 cm 15 20 cm 20 30 cm

30 40 cm

0% 10% 20% 30% 40% Relative 16S rRNA gene abundance of acidobacterial groups Figure 5. Relative abundance of acidobacterial subdivisions/groups in different depths of the Schlöppnerbrunnen II fen site in the years 2004 and 2007, as determined by of 16S rRNA gene targeted qPCR. Black circles indicate total numbers of bacterial and archaeal 16S rRNA genes. Relative abundances (given in % of the total number of bacterial and archaeal 16S rRNA genes) of acidobacterial groups 1, 2, and 3 are indicated in black, grey, and white bars, respectively. Error bars are the standard deviations of the mean for the three biological replicates.

42 Sulfate reducers in peatlands

Members of group 1 were most abundant, followed by members of group 2 and 3: Mean copy numbers per gram wet soil were 4.3 ± 1.3 x 106, 3.9 ± 3.6 x 106, and 2.4 ± 0.4 x 106 in the year 2004 and 9 ± 2.2 x 106, 5.6 ± 5 x 106, and 4.9 ± 0.9 x 106 in 2007 for group 1, group 2, and group 3, respectively. The relative abundance was 10.3% ± 5.1, 8.5% ± 1.9, and 5.0% ± 1 in the year 2004 and 20.0% ± 3.5, 12.1% ± 6.1, and 3.5% ± 1.7 in 2007 for the groups 1, 2, and 3, respectively. In contrast to Syntrophobacterrelated microorganisms (see above), 16S rRNA gene abundance of any of the three acidobacterial groups was not correlated with the dsrA abundance of any of the novel OTUs 1 and 2 (p=0.1230.875, Pearson coefficients 0.058 0.521).

Diversity and biogeography of dsrAB in geographically separated wetlands. In order to evaluate if the dsrABcontaining microorganisms in the Schlöppnerbrunnen fens are endemic or show a wider geographic distribution, a modified DGGE assay was applied for fingerprinting of dsrB in nine wetlands located in Austria, Italy and Germany (Table S1, Figure S1). In comparison to the original DGGE forward primer DSRp2060F (38), which covers only 29% of all sequences with the primer binding site in the dsrAB database, the newly developed DSR1728F primer mix has a significantly improved in silico coverage of 95% (Table S5). This modified dsrBDGGE analysis yielded 11 to 40 bands for the different wetland soil samples. The lowest and highest numbers of DGGE bands were detected in the Berndorf wet meadow soil and the Roßbrand II fen soil, respectively. With exception of the Roßbrand peats I and II, located in close distance (~1 km) to each other, the dsrBDGGE profiles between replicate soil cores were more similar to each other than dsrBDGGE profiles between different wetland sites (Figure 6). Furthermore, all precipitationfed bogs showed greater similarity in dsrBDGGE banding pattern to each other than to other wetland sites. PCA resulted in four principal components (PC), which explained 74.1% of the DGGEbased community composition data, whereby the first two ordination axes (Figure 6) explained 48.8% of the variation. Correlation analysis between principal components and environmental factors revealed that PC1 was strongly negatively correlated with pH (correlation coefficient of 0.9963), whereas PC2 was moderately negatively correlated with sulfate (correlation coefficient of 0.7286) and nitrate (correlation coefficient of 0.6834). Furthermore, the different peatland sites showed no major relationship based on their geographical position. In contrast, the measured environmental factors (pH, concentrations of sulfate and nitrate) explained significantly the differences in the dsrBDGGE profiles of the investigated wetlands (rho=0.568, p<0.001).

43 Part I

1.0 ABC Schallhof (F)

Rasner Möser (F) 0.5 ABC

Krähmoos (B)

PC 2 PC pH AC B 0.0 Roßbrand II (F) Schremser Hochmoor (B) BGroße A A Heide (B) AB C BC C A B C Roßbrand II (F) Roßbrand I (F) 0.5 ABC Berndorf (WM) Nitrate Sulfate

1.0 0.5 0.0 0.5 1.0 PC 1 Figure 6. Relationships between dsrBDGGEbased community structure data and environmental factors (pH, sulfate and nitrate concentrations) in eight, geographically separated wetlands were determined by principal component analysis. For each wetland the positions of the three biological replicates (sampling cores A, B, and C) in the ordination biplot is depicted. Wetland type is indicated: bog (B), fen (F), wet meadow (WM). Grey shading additionally highlights grouping of bogs.

For identification, 160 individual DGGE bands were extracted from the gel, reamplified and sequenced. Unambiguous dsrB sequences were obtained from 59 DGGE bands and grouped into 22 DsrBOTUs (Figure 7, Table S7). Only few OTUs were closely affiliated with representatives of characterized SRM. Ten OTUs, including six out of eleven previously defined Schlöppnerbrunnen DsrABOTUs (75), were present in at least four out of eight peatlands. Nine OTUs of these widely distributed OTUs were present at bog and fen sites, while dsrBDGGE OTU 6 was only detected in fens.

PCA of individual dsrB-DGGE bands closely related to Schlöppnerbrunnen dsrABOTUs (Table S7) indicate broad distribution of members of Schlöppnerbrunnen OTU 1 along the pH gradient and a preferential association with higher sulfate concentrations. In contrast, DGGE bands corresponding to Schlöppnerbrunnen OTUs 2, 3, and 6 were positioned at low pH and low sulfate concentrations in the PCA plot (data not shown).

44 Sulfate reducers in peatlands

Desulfobacteraceae 1 1 DG83 (F) Schlöppnerbrunnen II * Schlöppnerbrunnen OTU 5 * Everglades clone F1SU06, AY096050 (F) Rasner Möser Desulfomicrobium spp. (F) Roßbrand I Desulfonatronovibrio hydrogenovorans, AF418197 Desulfohalobium retbaense, AF418190 (F) Roßbrand II Desulfovibrio spp. (F) Schallhof Desulfovibrio spp. Desulfonatronum lacustre, AF418189 (B) Krähmoos Schlöppnerbrunnen soil clone, AM179486 Schlöppnerbrunnen OTU 9, AY167475 (B) Schremser Hochmoor Desulfomonile tiedjei, AF334595 (B) Grosse Heide Desulfotomaculum spp. Desulfobacterium anilini, AF482455 sulfate reducing strain mXyS1, AF482456 (WM) Berndorf Everglades clone F1SP02, AY096038 Everglades clone F1SP16, AY096042 Schlöppnerbrunnen soil clone, AM179488 Desulfobulbaceae Everglades clone F1SU18, AY096054 Thermodesulfobacterium spp. Syntrophobacter wolinii, AF418192 Schlöppnerbrunnen soil clone , AM179462 Schlöppnerbrunnen OTU 6 21 21 10% Other Syntrophobacteraceae * * Everglades clone U3SP20, AY096066 Everglades clone U3SP34, AY096068 Desulfacinum infernum, AF418194 Thermodesulforhabdus norvegica, AF334597 Desulfoarculus baarsii, AF334600 Schlöppnerbrunnen OTU 1 17 17 Schlöppnerbrunnen OTU 4 20 * *20 Schlöppnerbrunnen soil* clone, AM179502 * Everglades clone F1SU26, AY096056 2 DG154 2,3 *3 DG151 * Desulfobacca acetoxidans, AY167463 * Schlöppnerbrunnen OTU 2 18 18 Schlöppnerbrunnen soil clone, *AM179497 *4 DG13 Schlöppnerbrunnen soil clone, AM179455 4,5 Schlöppnerbrunnen soil clone, AM179464 *5 DG15,DG16, DG26 * Schlöppnerbrunnen soil clones, AM179495, AM179496 * Schlöppnerbrunnen OTU 7, AY167483 DG44,DG45,DG46,DG50 6 peatland clones New York 6 * Schlöppnerbrunnen clone, AM179478 * Everglades clones, AY096043, AY096052, AY096048 peatland clones New York, DQ855249, DQ855250, DQ855260, DQ855257, DQ855255, DQ855261 Schlöppnerbrunnen OTU 10, AY167474 peatland clone New York, DQ855252 Rasner Möser OTU 5 (Rm23) [1] Schlöppnerbrunnen soil clone , AM179453 Schlöppnerbrunnen soil clone, AM179473 Schlöppnerbrunnen soil clone, AM179482 Rasner Möser OTU 8 (Rm29) [1] 7 Schlöppnerbrunnen soil clone, AM179472 7 DG17 *8 Everglades clone U3SU24, AY096073 *8 mudflat clone, AY953402 DG129 * Wadden Sea sediment fosmid ws7f8, CT025836 * Schlöppnerbrunnen OTU 8, AY167476 peatland clones New York Schlöppnerbrunnen OTU 11, AY167472 9 22 Rasner Möser OTU 11 (Rm47) [1] *9 DG155 Everglades* clone F1SP13, AY096041 Everglades clone U3SP11, AY096062 * Everglades clone U3SU03, AY096069 Everglades clone U3SU27, AY096074 mudflat clone, AY953410 Rasner Möser OTU 7 (Rm27) [1] Rasner Möser OTU 10 (Rm71) [2] Schlöppnerbrunnen soil clone, AM179489 Rasner Möser OTU 6 (Rm26) [1] 16 16 Schlöppnerbrunnen soil clone, AM179506* 10 Schlöppnerbrunnen soil clone, AM179481 *10 DG144 * Rasner Möser OTU 3 (Rm9) [1] Schlöppnerbrunnen soil clone, AM179487 * deepsea sediment clone, AB263164 marine sediment clone, AM236163 Rasner Möser OTU 4 (Rm22) [1] deepsea sediment clone, AB263161 peatland clone New York, DQ855256 Rasner Möser OTU 12 (Rm58) [1] Thermosinus carboxydivorans Nor1, NZ_AAWL01000006 Pelotomaculum sp. MGP, AB154391 11 Desulfitobacterium spp. 11 DG158, DG159 * Desulfosporosinus orientis, AF271767 * Everglades clone U3SP09, AY096061 Carboxydothermus hydrogenoformans Z2901, NC_007503 Desulfotomaculum spp. Thermodesulfobium narugense, AB077818 13 Everglades clone U3SP30, AY096067 13 DG33 groundwater clone, EF065023 * Rasner Möser OTU 9 (Rm72) [4] * Schlöppnerbrunnen soil clone, AM179484 groundwater clone, EF065095 Rasner Möser OTU 2 (Rm25) [2] Schlöppnerbrunnen OTU 3 Rasner Möser OTU 1 (Rm19) [20] 19 19 12 * *12 DG146 * peatland clones New York * 14 groundwater clone, EF065069 14 DG64, DG66, DG87 *15 *15 DG116 * Archaeoglobus * Thermodesulfovibrio spp.

45 Part I

Figure 7. DsrAB consensus tree showing the affiliation of dsrAB clone and dsrB DGGE sequences recovered nine different wetlands; five fens (F), three bogs (B), and a wet meadow (WM). OTUs from the Schlöppnerbrunnen (75) and Rasner Möser fens were based on DsrAB sequences longer than 500 amino acids and are depicted in bold and colored type. For each Rasner Möser OTU, a representative clone and the number of sequenced clones are given in round and square brackets, respectively. Sequences shorter than 500 amino acids are indicated by dashed branches and asterisks, the latter depict the phylogenetic positions of 22 dsrBOTUs, which were identified by sequencing of dsrBDGGE bands. In addition, the presence/absence of each dsrBOTU in the nine wetlands was inferred from comparison of DGGE banding pattern and is indicated by presence/absence of the respective colored square. Scale bar indicates 10% estimated sequence divergence.

Soil samples from the groundwaterfed fen site Rasner Möser were additionally subjected to dsrAB clone library analysis. A total of 39 fully sequenced dsrAB clones (~1900 bp) formed 12 OTUs; resulting in a Good’s coverage of 78%. Only OTU 12 was moderately related to Thermosinus carboxydovorans, while the remaining OTUs had no close cultivated relative (Figure 7). Rasner Möser OTUs 1 (represented by more than half of the dsrAB sequences, n=20) and 11 were most closely related to the deepbranching Schlöppnerbrunnen OTUs 3 (88.297.3% amino acid identities) and 11 (90% amino acid identity), respectively. Only these two Rasner Möser OTUs and OTU 6 were also identified by dsrBDGGE analysis (Table S7).

Discussion

Novel lineages are ‘core’ organisms and dominate a temporally stable but spatially heterogeneous dsrABcontaining population in the Schlöppnerbrunnen II fen.

Environmental diversity surveys of the dissimilatory (bi)sulfite reductase genes dsrAB have repeatedly demonstrated that clone libraries from diverse ecosystems can be largely composed of sequences forming new branches with no cultivated representative in the DsrAB tree (66, 75). These novel dsrAB sequences can be indicative of sulfite/sulfate reducers (or organosulfonate reducers) that belong to either previously undescribed phylogenetic lineages or described lineages not yet known to contain these functional guilds of microorganisms.

46 Sulfate reducers in peatlands

Additionally, anaerobes that live in syntrophic association with other microorganisms might be carriers of these novel dsrAB, which are presumably genomic remains of an ancestral sulfate reducing metabolism (50, 120). The fact that evolutionary distribution of dsrAB among SRM was influenced by some lateral gene transfer events (59, 129) complicates phylogenetic interpretation of environmentally retrieved dsrAB sequences. Hence, these novel dsrAB sequences might also derive from microorganisms which are phylogenetically related to known cultivated microorganisms but have received their dsrAB via lateral gene transfer from a yet unknown donor. The true identity and also the environmental relevance of microorganisms carrying these novel dsrAB types are currently unknown.

Here, we have developed and extensively tested the diagnostic performance of a habitat specific dsrABbased microarray and three OTUspecific qPCR assays for revealing the dynamics of the dsrABcontaining microbial community in the Schlöppnerbrunnen II fen, a model peatland and longterm study site that was previously shown to harbor several novel dsrAB lineages (75, 106). As expected, qPCR assays were more sensitive than FGA analysis (Figure 1A, Figure 2). However, qPCR results confirmed the experimentally inferred detection limit of the FGA (0.01% of the total microbial community) as dsrABOTUs with abundances of ≥0.05% (relative to the total number of bacterial and archaeal 16S rRNA genes) were also consistently detected in fen soil samples by microarray analysis. FGA hybridization pattern of soils sampled over a period of six years indicated little longterm changes in the number and types of dsrABcontaining microorganisms, confirming that members of several dsrABlineages are part of the autochthonous community in the Schlöppnerbrunnen II fen. In contrast, clear spatial differences were evident in FGA hybridization pattern, with only very few probetarget groups being detected in the most upper fen layer (Figure 1). Consistent with this observation, copy numbers of dsrA of OTUs 1, 2, and 6 analyzed by qPCR were always considerably lower in the most upper than in deeper soil layers. Interestingly, the overall dynamics of the dsrAB harboring community are mirrored by spatially highly variable but temporally relatively stable sulfate concentrations at the study site (2). In contrast, highly variable δ34S values were indicative of past changes in the activity of fen soil SRM in space and time. Comparably stable but low bulk concentrations of sulfate over time thus rather reflect an active buffering/recharging system for sulfate than balanced consumption and production of sulfate at the Schlöppnerbrunnen II fen site (2). Factors other than sulfate availability must thus be responsible for the reduced dsrAB diversity in the most upper soil layer. While generally regarded as anaerobic microorganisms, it is well known that some SRM (i) can be highly active

47 Part I in oxicanoxic transition zones and even oxic environments (84, 85, 113) and (ii) tolerate or respire oxygen in pure culture (22, 33, 68, 108). However, to date growth under oxic conditions was shown only for one sulfate reducing bacterium, Desulfovibrio desulfuricans ATCC 27774 (72). Recurring events of drought and precipitation are major determinants of short and long term changes in the water table and thus the redox status of peatlands (28, 99). Oxygen exposure in upper peat zones is more substantial than in deeper zones due to increased impact of drought events (28, 99) and release of oxygen from aerenchymatic plant roots (123). It is thus conceivable that growth inhibition due to prolonged exposure to oxygen restricts the abundance of anaerobic, dsrABcontaining microorganisms in the upper fen soil layer (approximately 010 cm) on a longterm scale. Interestingly, FGA and qPCR analyses both indicated that members of novel dsrAB lineages (e.g. OTUs 1, 2, 3, 4, and 11) outnumbered members of lineages that contain cultivated microorganisms. In particular, all soil fractions contained a higher copy number of dsrA from the novel OTUs 1 and 2 than from the Syntrophobacter woliniirelated OTU 6. In some soil samples from depths of 1030 cm, members of OTUs 1 and 2 even accounted for a significant proportion of the total microbial community (12% dsrA copy number relative to the total number of bacterial and archaeal 16S rRNA genes) (Figure 2). These relatively high abundances of ≥0.1 1% imply that unknown OTU 1 and OTU 2 members are socalled ‘core’ microorganisms (93) and actively contribute, at least occasionally, to the prevalent biogeochemical processes in the fen.

Identity and putative functional properties of dominant Schlöppnerbrunnen II fen microorganisms.

Classical 16S rRNA gene clone library analysis showed that common community members in the model fen belong to twelve bacterial and two archaeal phyla. The list of 85 bacterial and 18 archaeal specieslevel 16S rRNAOTUs detected in the fen represents a first general, albeit incomplete inventory of microorganisms that might harbor the novel dsrAB gene types. Clones from Acidobacteria groups/subdivisions 1, 2, and 3 (7, 47, 128) accounted in total for over 50% of the bacterial 16S rRNA gene sequences. Although acidobacterial sequences tend to be overrepresented in general clone libraries [e.g. (26, 60)], qPCRbased quantification of the three main acidobacterial groups supported their high frequency of occurrence in the fen soil (5.6 26.3%, 615.5%, and 3.613.1%, for group 1, 2, and 3, respectively) and comparably stable distribution across soil layers (Figure 5). High abundance of these microorganisms in deeper, mainly anoxic peat layers implies an anaerobic or facultative aerobic lifestyle of at least some

48 Sulfate reducers in peatlands group members. No correlation was found between abundances of individual acidobacterial groups and dsrABOTUs 1 or 2. This could have two reasons: First, members of these Acidobacteria subdivisions in the peat are not carriers of these novel dsrAB types. Second, because qPCR assays target microorganisms at highly different phylogenetic levels (subdivision level for Acidobacteria versus approximately species level for dsrABOTUs), it is possible that some acidobacterial group members harbor these dsrAB but correlations are obscured by other group members lacking these genes. Independent from being dsrAB carriers or not, Acidobacteria are commonly detected in other acidic peatlands and akin environments (6, 26, 52) and thus presumably have an important ecological role in such soil ecosystems. It is likely that the phylogenetically different acidobacterial OTUs (n=33) in the Schlöppnerbrunnen II fen also have different physiological features. Assigning general metabolic capabilities across all detected members of the entire phylum is difficult, if not impossible. Nevertheless, the following physiological traits have been attributed to Acidobacteria and provide some hints why they are successful inhabitants of acidic peatlands. First, many isolated representatives are moderately acidophilic or acid tolerant and favor environments with pH 3.5 to 6.5 (57, 65, 90, 91). This is especially true for members of subdivision 1 (104), the most abundant group detected in our study (Figure 4). Second, peat soil Acidobacteria are able to utilize a variety of carbon substrates, in particular complex, plant derived compounds (90, 91, 122). 16S rRNAbased stable isotope probing of the Schlöppnerbrunnen fen has indicated that some members of group 1 (clone AM773995) are involved in fermentation of the monosaccharide xylose, an important ligninderived intermediate in the carbon degradation network in wetlands (43). Furthermore, aerobic strains of the species Bryobacter aggregatus (subdivision 3), isolated from acidic Sphagnum peatlands, consumed galacturonic and glucuronic acids, which are typically released from the cell walls of decaying Spagnum mosses (65). Third, genomic and physiological data postulate that ecological functions of some Acidobacteria are nitrate/nitrite reduction and iron oxidation/reduction (91, 122);(90), redox processes that are also taking place in the Schlöppnerbrunnen fen model system (2, 64, 101).

Most of the detected bacterial OTUs represent novel family/genuslevel phylotypes within known phyla, while only eight OTUs could be assigned to described genera. Interestingly, members of four of those genera, namely Smithella, Syntrophobacter, Desulforhopalus, and Desulfosporosinus, are endowed with metabolic features for sulfate reduction and/or syntrophic growth. The propionateoxidizing syntrophs Smithella propionica and Syntrophobacter wolinii

49 Part I were both initially cultivated in coculture with hydrogen/formateutilizing methanogens (71). Potential methanogenic partners for a thermodynamically favorable syntrophic lifestyle have been detected in the fen and belong to the obligate acetateconsuming family Methanosaetaceae and the hydrogen/formateconsuming family Methanomicrobiaceae (Figure 4) (44, 125). Smithella- and Syntrophobacterrelated bacteria were actively involved in syntrophic oxidation of propionate in a flooded rice field soil and it is tempting to speculate that the Schlöppnerbrunnen II microorganisms have the same function. Although propionate is an important carbon degradation intermediate, which is rapidly formed and consumed after anoxic incubation of fen soil, DNA stable isotope probing of Schlöppnerbrunnen II soil that was incubated for two months with a 13Clabelled mixture of acetate, propionate, formate, and lactate showed no indications for 13Clabeling of Smithella- or SyntrophobacterDNA (Pester et al., submitted). However, growth of these bacteria under thermodynamically limited, syntrophic conditions was probably too low to sustain sufficient labeling of their DNA for separation by density gradient centrifugation (79). One fen OTU has specieslevel 16S rRNA sequence identity to Desulforhopalus vacuolatus, a deltaproteobacterial SRM that can use sulfate, sulfite or thiosulfate as electron acceptor for incomplete oxidation of propionate, lactate, pyruvate, propanol or ethanol (51). Additionally, disproportionation of sulfite allows D. vacuolatus to grow on acetate as sole carbon source. In the absence of an electron acceptor, D. vacuolatus is able to switch to fermentation of lactate or pyruvate. Although Desulforhopalus and Syntrophobacter species are capable of dissimilatory sulfate reduction, only Desulfosporosinus species were shown to be catalyzing this process in the Schlöppnerbrunnen II fen (Pester et al., submitted). Albeit Desulfosporosinus species are members of the ‘rare biosphere’ (<0.01% of the total archaeal and bacterial community) in the fen, they can sustain such a high metabolic activity to account for a major part of the bulk sulfate reduction rates measured in situ. However, besides sulfate also thiosulfate or sulfite, present in high amounts in the fen system (200.3 mg/kg S) (96), might be employed by the detected SRM as alternative electron acceptors for anaerobic growth. It is unlikely that some of the novel dsrAB types derive from the genera Syntrophobacter, Desulforhopalus, and Desulfosporosinus, because dsrAB sequences related to sequences from cultivated members of these genera have already been recovered from the Schlöppnerbrunnen fens (9, 75) (Pester et al., submitted). However, archaeal sulfite reducers of the genus Pyrobaculum contain two or more very distantly related copies of dsrAB in their genomes (55 91% DsrAB sequence identity) (4, 34). Although the genome of Syntrophobacter fumaroxidans

50 Sulfate reducers in peatlands

MPOB has only one copy of dsrAB (http://genome.jgipsf.org/synfu/synfu.home.html), it is at least theoretically conceivable that some of the Syntrophobacter, Desulforhopalus or Desulfosporosinus fen species in the fens contain in addition to their orthologous dsrAB a xenologous dsrAB copy that was acquired from a yet unknown donor via lateral gene transfer. Although members of the genus Smithella are not capable of dissimilatory sulfite/sulfate reduction, they could still harbor dsrAB, analogous to the situation in other syntrophs such as some Pelotomaculum species (48, 49), but this remains to be shown.

Novel dsrABcarrying microorganisms are widespread in wetlands.

DGGE analyses of nine wetlands with geographical distances ranging from 1 km to 400 km showed that many of the dsrABcontaining microorganisms (six OTUs) found in the Schlöppnerbrunnen fens are not endemic but also inhabit other peatlands. Previous molecular fingerprinting studies of anthropogenically impacted sediments (94) and river floodplains (83) observed signatures of biogeography in dsrAB data, and the overall dsrBDGGE banding pattern also indicated that the spatial distribution of dsrABcontaining microorganisms between the nine wetland sites is nonrandom. On condition that dsrBDGGE analysis with degenerated primers is prone to biases [e.g. (11, 87, 88)] and restricted by its detection limit, we sought to identify environmental and/or geographic factors that govern the biogeography of dsrAB containing microorganisms in wetlands. Spatial variations in microbial community composition can generally depend on contemporary environmental factors, on past events (such as dispersal limitation and past environmental conditions) that lead to genetic differentiation and possibly speciation, or on both (82). If community structure is only influenced by the currently prevailing environmental parameters, known as BaasBecking hypothesis “everything is everywhere, but the environment selects” (25), we would expect no geographical distance dependent differences. Indeed, geographical distance between sites was found to be a poor predictor of dsrBDGGE banding pattern distribution, suggesting that the current biogeography of dsrABcontaining microorganisms in wetlands is not a legacy of historical events (82). Although only few environmental parameters (Table S1) were measured, the observed biogeography pattern significantly reflected the influence of environmental variations. In particular, soil pH was the dominant factor that determined the community structure. This finding is consistent with a general study of bacterial biogeography in soils, where bacterial richness, diversity, and overall community composition were found to be mainly influenced by soil pH (32). In our study, the dsrABcontaining communities in the wetlands seemed to be additionally

51 Part I impacted by the source of water, as bogs, which are fed by precipitation, clustered together in principal component analysis (Figure 6). The type of water source can be regarded as a for the nutrient content of peatlands (12, 62). Therefore, microorganisms inhabiting precipitationdependent bogs generally have to cope with a lower availability of minerals and nutrients compared to microorganisms that live in fens, which receive additional substrates by the groundwater flow. Phylogenetic analyses of dsrBDGGE bands and a dsrAB clone library from the Rasner Möser fen revealed ten OTUs that were broadly distributed among different bogs and fens; these OTUs include the Syntrophobacter woliniirelated OTU and nine OTUs with no close cultivated relatives. However, only microorganisms belonging to one of these more widely distributed OTUs (dsrBDGGE OTU 6) appeared to preferentially colonize fens (Figure 7) and thus could be potential bioindicators for this type of wetland.

Conclusions

We show that microorganisms with phylogenetically novel types of the dissimilatory (bi)sulfite reductase genes dsrAB are persistent and numerically significant inhabitants of a longterm experimental peatland field site. Comparative analysis of dsrABcarrying microorganisms in different peatlands has identified soil pH as one of the most influential community structure shaping environmental factors. In contrast, dispersal limitation of dsrABcarrying microorganisms does not seem to contribute largely to endemism, because members of various novel dsrAB lineages display a rather cosmopolitan distribution among peatlands that are located up to several hundred kilometers apart. Numerical abundance and broad distribution of these novel dsrABcarrying microorganisms clearly suggest that they have a considerable impact on peatland ecosystem functioning. Revealing the identity and actual physiological features of these mysterious microbes thus far proved to be a challenge (Pester et al., submitted). However, the relatively high abundance of these yet unidentified microorganisms at a given time and location holds much promise for future research, because it makes them amenable to techniques that allow sorting and genomic or physiological analysis of individual microbial cells (45, 70, 110, 126).

52 Sulfate reducers in peatlands

Acknowledgements

We thank Stephan Duller for help with dsrA primer design/evaluation, Achim Schmalenberger for providing dsrAB clones for microarray evaluation, and Christian Baranyi for help with statistics and excellent technical assistance. We are grateful to Jen How Huang, Janine Franke, Alexandra Hamberger, Marco Reiche, and Florian Wenter for their help with sampling. We thank Andreas Richter and Christina Kaiser for ionchromatographic determination of anion concentrations. This research was financially supported by the Austrian Science Fund (P18836 B17, AL), the Alexander von HumboldtFoundation (MP), the German Federal Ministry of Science and Education (BIOLOG/BIOTA project 01LC0621D; MW and AL), the Austrian Federal Ministry of Science and Research (ASEAUNINET; PD) and the University of Vienna (PD).

53 Part I

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63 Part I

Supplementary material

Figure S1. Geographic location of the investigated wetlands.

64 Sulfate reducers in peatlands

Desulfobacteraceae Schlöppnerbrunnen I soil clone dsrSbI50, AY167464 Schlöppnerbrunnen soil clone, AM179464 41 Schlöppnerbrunnen I soil clone dsrSbI82, AY167482 Schlöppnerbrunnen soil clone, AM179472 30 40 Desulfohalobium retbaense, AF418190 Schlöppnerbrunnen soil clone, AM179457 56 Desulfomicrobium spp. Schlöppnerbrunnen I soil clone dsrSbI64, AY167474 57 Schlöppnerbrunnen soil clone, AM179501 56 Desulfonema limicola, AY626031 Schlöppnerbrunnen soil clone, AM179473 52 Desulfovibrio burkinensis, AB061536 Schlöppnerbrunnen soil clone, AM179478 58 Desulfovibrio cuneatus, AB061537 Schlöppnerbrunnen soil clone, AM179453 SbI enrichment clone 115 27 Schlöppnerbrunnen soil clone, AM179468 51 Desulfovibrio vulgaris, U16723 Schlöppnerbrunnen soil clone, AM179480 54 Schlöppnerbrunnen soil clone, AM179458 Desulfovibrio spp. Schlöppnerbrunnen soil clone, AM179482 Desulfonatronum lacustre, AF418189 Schlöppnerbrunnen soil clone, AM179510 55 53 Schlöppnerbrunnen soil clone, AM179486 28 Schlöppnerbrunnen soil clone, AM179471 Desulfomonile tiedjei, AF334595 Schlöppnerbrunnen soil clone, AM179493 Schlöppnerbrunnen I soil clone dsrSbI88, AY16747529 Schlöppnerbrunnen soil clone, AM179496 42 Schlöppnerbrunnen soil clone, AM179452 Schlöppnerbrunnen soil clone, AM179483 44 Schlöppnerbrunnen soil clone, AM179488 17 Schlöppnerbrunnen soil clone, AM179495 43 Schlöppnerbrunnen soil clone, AM179469 19 Schlöppnerbrunnen II soil clone dsrSbII2, AY167483 50 Schlöppnerbrunnen soil clone, AM179485 18 20 Schlöppnerbrunnen soil clone, AM179477 47 Schlöppnerbrunnen soil clone, AM179455 46 45 Firmicutes spp. Schlöppnerbrunnen soil clone, AM179497 48 Schlöppnerbrunnen II soil clone dsrSbII3, AY167467 Desulfobulbaceae Schlöppnerbrunnen II soil clone dsrSbII28, AY167479 Schlöppnerbrunnen II soil clone dsrSbII34, AY167468 Desulfotalea spp. 49 Desulforhopalus singaporensis, AF418196 Archaeoglobus spp. SbI enrichment clone 2110 Schlöppnerbrunnen II soil clone dsrSbII25, AY167481 SbI enrichment clone 311 Schlöppnerbrunnen II soil clone dsrSbII36, AY16746932 SbI enrichment clone 3110 26 Schlöppnerbrunnen soil clone, AM179466 31 Thermodesulfobacterium spp. Schlöppnerbrunnen soil clone, AM179456 33 Schlöppnerbrunnen soil clone, AM179484 34 Syntrophobacteraceae Firmicutes spp. Syntrophobacter wolinii, AF418192 Schlöppnerbrunnen soil clone, AM179462 24 Schlöppnerbrunnen soil clone, AM17948735 Schlöppnerbrunnen soil clone, AM179459 23 21 Carboxydothermus hydrogenoformans, NC_007503 Schlöppnerbrunnen II soil clone dsrSbII40, AY167465 Schlöppnerbrunnen soil clone, AM179489 37 Schlöppnerbrunnen I soil clone dsrSbI54, AY167478 Schlöppnerbrunnen soil clone, AM179481 36 Schlöppnerbrunnen soil clone, AM179461 Schlöppnerbrunnen soil clone, AM179451 38 Schlöppnerbrunnen soil clone, AM179470 22 25 Schlöppnerbrunnen soil clone, AM179475 Desulfoarculus baarsii, AF334600 Schlöppnerbrunnen soil clone, AM179504 Schlöppnerbrunnen soil clone, AM179492 Schlöppnerbrunnen soil clone, AM179494 Schlöppnerbrunnen soil clone, AM179506 39 Schlöppnerbrunnen soil clone, AM179476 Schlöppnerbrunnen I soil clone dsrSbI75, AY167476 16 Thermodesulfovibrio islandicus, AF334599 Schlöppnerbrunnen soil clone, AM179460 13 Thermodesulfovibrio yellowstonii, U58122, U58123 Schlöppnerbrunnen soil clone, AM179474 14 Schlöppnerbrunnen II soil clone dsrSbII39, AY167472 15 10% Schlöppnerbrunnen soil clone, AM179500 Schlöppnerbrunnen soil clone, AM179509 Schlöppnerbrunnen soil clone, AM179502 Schlöppnerbrunnen soil clone, AM179499 Schlöppnerbrunnen soil clone, AM179503 11 Schlöppnerbrunnen soil clone, AM179463 12 10 Desulfobacca acetoxidans, AY167463 Schlöppnerbrunnen II soil clone dsrSbII15, AY167471 Schlöppnerbrunnen II soil clone dsrSbII9, AY167477 2 9 Schlöppnerbrunnen soil clone, AM179498 5 3 Schlöppnerbrunnen soil clone, AM179490 4 Schlöppnerbrunnen soil clone, AM179465 5 7 8 Schlöppnerbrunnen soil clone, AM179450 2 6 Schlöppnerbrunnen soil clone, AM179467 Schlöppnerbrunnen soil clone, AM179479 Schlöppnerbrunnen I soil clone dsrSbI60, AY167466 Schlöppnerbrunnen I soil clone dsrSbI66, AY167480 Schlöppnerbrunnen II soil clone dsrSbII20, AY167470 Schlöppnerbrunnen soil clone, AM179454 Schlöppnerbrunnen soil clone, AM179491 Schlöppnerbrunnen soil clone, AM179505 Schlöppnerbrunnen soil clone, AM179507 Schlöppnerbrunnen soil clone, AM179508 Schlöppnerbrunnen I soil clone dsrSbI59, AY167473 1

Figure S2. Target groups of dsrABbased oligonucleotide microarray probes. DsrAB phylogeny of Schlöppnerbrunnen dsrAB sequences (6, 7) was inferred by distance matrix analysis. Colored boxes indicate perfectlymatched target sequences of probetarget groups 1 to 58, as defined in Table S4.

65 Part I 8 9 7 1 2 3 4 5 6 11 10 14 15 17 18 20 21 24 28 29 31 34 35 39 40 43 46 48 49 52 55 56 57 13 16 19 22 23 25 26 27 30 32 33 36 37 38 41 42 44 45 47 50 51 53 54 58 2001 12

0 – 10 cm > 0.39 0.36 0.39 0.33 0.36 10 – 17.5 cm 0.30 0.33 0.27 0.30 0.24 0.27 17.5 – 25 cm 0.21 0.24 0.18 0.21 0.12 0.18 0.12 0.15 25 – 30 cm 0.09 0.12 0.06 0.09 0.03 0.06 2004 < 0.03

0 – 5 cm

5 – 10 cm

10 – 15 cm

15 – 20 cm

2007

0 – 5 cm

5 – 10 cm

10 – 15 cm

15 – 20 cm

20 – 30 cm

30 – 40 cm

Figure S3. Microarraybased dsrAB diversity analysis of Schlöppnerbrunnen II fen soils sampled from different depths in the years 2001, 2004, and 2007. Results of microarray hybridizations are displayed as mean nSBR of all probes within a probetarget group and are shown for each replicate. The color code translates into different mean nSBR values. Probe target groups are arranged phylogenetically according to their position in a dsrAB neighbor joining tree.

66 Sulfate reducers in peatlands

TableS1. Description and location of sampling sites.

Geographical a Sulfate Nitrate Wetland name Wetland type pH c c position [M] [M] 4.0 Schlöppnerbrunnen 50° 09' 21'' N fen 6.1b 22240 b <5120b 11° 52' 22'' O Germany Krähmoos 46° 47' 60'' N bog 4.0 10.2 1.1 11° 54' 18'' O Italy Rasner Möser 46° 48' 27'' N fen 4.3 9.1 0.2 12° 04' 25'' O Italy Roßbrand I 47° 24' 27'' N fen 4.1 18.3 2.1 13° 28' 27'' O Austria Roßbrand II 47° 24' 39'' N fen 4.9 d 57.5 d 2.3 d 13° 28' 56'' O Austria Große Heide 48° 34' 06'' N bog 5.3 41.1 12.3 14° 45' 26'' O Austria Schremser Hochmoor 48° 48' 17'' N bog 4.7 30.5 1.8 15° 06' 04'' O Austria Schallhof 47° 53' 23'' N fen 7.3 2.1 0.2 15° 56' 43'' O Austria

Berndorf 47° 56' 41'' N wet meadow 7.6 116.8 0.5 16° 06' 46'' O Austria a Wetland classification: bog, ombrotrophic (rainwaterfed); fen, minerotrophic (mostly groundwaterfed) b Data from (7) c Sulfate and nitrate was determined in the soil water by ion chromatography d Soil core C was measured separately: pH = 5.8, nitrate: 3.3 M, sulfate: 95.6 M

67 Part I

Table S2. dsrAB-targeted primers.

Theoretical melting Primer Sequence (5’3’) G+C (%) a Reference temperature

DSR1F ACSCACTGGAAGCACG 63 4251 (9) DSR1Fa ACCCAYTGGAAACACG 5056 4345 (6) DSR1Fb GGCCACTGGAAGCACG 69 51 (6) DSR1Fc ACCCATTGGAAACATG 44 40 (10) DSR1Fd ACTCACTGGAAGCACG 56 47 (10) DSR1Fe GTTCACTGGAAACACG 50 42 Pester et al. submitted DSR1Ff AGCCACTGGAAACACG 56 47 Pester et al. submitted DSR1Fg GGCCACTGGAAACATG 56 44 Pester et al. submitted DSR1Fh GGCTATTGGAAGCACG 56 46 Pester et al. submitted DSR4R GTGTAGCAGTTACCGCA 53 48 (9) DSR4Ra GTGTAACAGTTTCCACA 41 41 (6) DSR4Rb GTGTAACAGTTACCGCA 47 45 (6) DSR4Rc GTGTAGCAGTTKCCGCA 5359 49 (6) DSR4Rd GTGTAGCAGTTACCACA 47 43 (10) DSR4Re GTGTAACAGTTACCACA 41 40 (10) DSR4Rf GTATAGCARTTGCCGCA 4753 4748 Pester et al. submitted DSR4Rg GTGAAGCAGTTGCCGCA 59 52 Pester et al. submitted a Melting temperatures were estimated using oligoCalc (5)

Table S3. Primers and targets for PCR and in vitro transcriptionbased preparation of a defined mixture of fluorescentlylabelled control RNA for microarray hybridization.

PCR Amount of annealing labeled target Target Primer Primer sequence (5’3’) Reference temp. RNA per (°C) hybridization

16S rRNA gene of Desulfonema BACT8F AGAGTTTGATYMTGGCTC (4) 52 100 ng limicola DSM 2076 BACT1529R CAKAAAGGAGGTGATCC

Nucleotide transporter NTT1 gene of NTT1F ATGTCGCAAGATGCGAAA (8) 55 75 ng Protochlamydia amoebophila UWE25 NTT1R TTAGCTAGTAGCTATTTC

16S rRNA gene of Desulfohalobium BACT8F AGAGTTTGATYMTGGCTC (4) 52 50 ng retbaense DSM 5692 BACT1529R CAKAAAGGAGGTGATCC

Nucleotide transporter NTT5 gene of NTT5F ATGAAAAATCAACAAAAT (3) 55 25 ng Protochlamydia amoebophila UWE25 NTT5R TTATCCATGGGAAGCTTC

Nucleotide transporter NTT4 gene of NTT4F ATGAGTAAAACAAACCAG (2) 55 10 ng Protochlamydia amoebophila UWE25 NTT4R TTATTTTTTTATAAAAGC

68 Sulfate reducers in peatlands

Table S4. Oligonucleotide probes for microarray hybridization.

Probe Perfectlymatching target Target dsrAB deltaG Probe Probe name target d Sequence (5'3') organisms/sequences gene OTU [kcal/mol] length group dsrCONT1 Desulfohalobium retbaense 16S rRNA CGCAGAGTTATCCCATACCTCGAGGTAGATTATC 41.2 34 dsrCONT2 DSM 5692 T 16S rRNA GACCCGTATTGGGAATCACCCATTTCTTCCCT 40.8 32 dsrCONT3 Desulfonema limicola DSM 16S rRNA GCTCCCCGAAGGGCACTATCTCCTTTCAAAG 40.7 31 dsrCONT4 2076 16S rRNA TAGAGGCCATCTTTCATCATTAATACCGGGGTA 40.7 33 dsrCONT7 Protochlamydia amoebophila ntt1 GGATAGCTGCAGGGACAACACCAAAAGACTTC 41 32 dsrCONT8 UWE25 ntt1 GCCATGCATCAACATTTGCAGGTAAATGCTTG 40.6 32 dsrCONT9 Protochlamydia amoebophila ntt4 GCGTGATTATAGGTCCCATTACAGCGCCTAAA 40.5 32 dsrCONT10 UWE25 ntt4 ATCCATATTGGCTGCAAAACGCGTTTGTGCAA 42.2 32 dsrCONT11 Protochlamydia amoebophila ntt5 GCCATATTTCCAACGAACATACAGAGGGGGTA 40.5 32 dsrCONT12 UWE25 ntt5 TGTCAGTCCAAACCCCACTACCAATACAGCTA 40.7 32 dsrA1 dsrA TCAGTTCGGCGAAGGTGGGTTCCAGTTCAT 40.8 30 dsrA2 AY167466, AY167480, dsrA AAGGTGGGTTCCAGTTCATCGGTGGTGGTG 40.7 30 AM179479, AM179467, dsrA3 AY167470, AY167473, dsrA TTCATCGGTGGTGGTGCCCAGGAAGATCAT 40.1 30 1 OTU 1 dsrB4 AM179507, AM179505, dsrB GTTGTCTTTGATGATGGGCGGCAAGAACTTGT 41.4 32 AM179491, AM179508, dsrA5 AM179454 dsrA CCAGGATGGGGGCGTGCGATCCCAGGAG 41.5 28 dsrB6 dsrB GGGGGGCCAATATCAGTTATACGGTCAGCC 39.6 30 dsrA7 dsrA ACGGGTTATCTCTGGGTGCCGAGACCATTT 40,0 30 dsrA8 dsrA GCACGCGCGATGGAAGCGACGCAATCG 41.6 27 dsrB9 AY167467, AY167479, dsrB CTCGCAAACTCGGGGAAGAATGTCGTGCTT 40.7 30 49 OTU 2 dsrB10 AY167468 dsrB ACGCCGGGCGCCAGGTTCTCGTGATACTT 42.4 29 dsrA11 dsrA GGGACAGCCCGCGCACTTGATCTTGAACTT 41.7 30 dsrB12 dsrB AGTTCTTCTTCACAATCGGAGGCAGGAACTTGT 41.8 33 dsrA13a dsrA CAATCATTGGGACAACCGGCACACTTGATTTT 40.5 32 dsrA14a AM179497 dsrA 48 GCTGGTATAAAACCATCCCGCCGGATGGTT 40,0 30 dsrA15a dsrA GTGCATGTTGGTCAATCCGGATCCGTGCTT 40.8 30 dsrA16 dsrA TGGAAAGCCGGCTCAAGTTCCGGTGTGGTT 42.1 30 dsrA17 AM179495 dsrA 43 GCCCAAATCGAATCCCGCCCTCGCAAGTT 41.1 29 dsrA18 dsrA GCGCCATACCGACGCAACACGAAGGCGT 42.2 28 dsrA19 dsrA TCGTAGGGCAGAGATCGCAAACGTGCTTTT 40.5 30 dsrB20 AY167483 dsrB 50 OTU 7 TACTCGCCGGATCTGGTCCTTTTGGACCTT 40.5 30 dsrA21 dsrA CACGCAGTTCCGGTTGTCAATCGCGAGTTT 41.5 30 dsrA22 dsrA CTCGAGTAGAACCACGCCGCCGGTTGGTT 41.8 29 AM179496, AM179471, dsrA23 dsrA 42 CATCGGGAAATTGCTCGCCTACGTCACTGT 40.4 30 AM179493 dsrA24 dsrA TGAAAGGTGGGCTCCAACTGGTCCGTGGTT 41.5 30 dsrA25 dsrA CGGCGTATTCGGCGACCTTCTCATCGTTTT 41,0 30 dsrA26 AM179468, AM179453 dsrA 51 CACGGATGGTGTGGAAGCGCGCCACGTT 42,0 28 dsrA27 dsrA GTGGAAGCGCGCCACGTTGGGGAATTCTT 41.4 29 dsrA28 dsrA CGCCTTGGCAAACTCCTGAAACGCGGGTT 41.5 29 dsrA29 AM179483 dsrA 44 GTGCACGTTGGTCACGCCCGAACCGTTG 41.5 28 dsrA30 dsrA CGCAGATGGTGCGAATGGCGTCACTGGTG 41.9 29 AM179510, AM179482, dsrA31 dsrA 53 CTCCTTGAAGGTGGGCTCGAGCTCATCGG 40.2 29 AM179458, AM179480 dsrA32 AM179510, AM179482, dsrA TCGGGCATCCACTGCACTTGAACTTGAACTT 41,0 31 55 dsrA33 AM179458 dsrA TTGACACGCAGCGTATGGAAGTGCGCGAC 41.8 29

69 Part I

dsrA34 dsrA CTCATCCTGGTAGGTCCGCGTCAAGTCGTC 41.1 30 dsrA35 dsrA GAGGTCGAATCCCGCGGCGGTCAGCTCC 42.3 28 AM179480 54 dsrA36 dsrA CTCATCCTGGTAGCTGCGCGTCAAGTCGTC 41.9 30 dsrA37 dsrA GCGGTCGCACACGTCTTTCTTGATGTCGAG 41.6 30 AM179501, AM179457 56 dsrA38 dsrA GACAGCGGTCGCACACGTCTTTCTTGATGT 40.9 30 dsrA39 dsrA ATACTCATCCCAGAATTCCCACACGCGTTTC 39.8 31 dsrB40b AY167474 dsrB 57 OTU10 CCGCAGCTCCTGCTTGAGCGGTTCGATCT 41.8 29 dsrA41 dsrA GCGTGGGGCAGCGATCGCACACATCCTT 41.6 28 dsrA42 dsrA CTTGAAGGCAGGCTCCAGCTCAGAGGTCTT 40.3 30 dsrA43 AM179478 dsrA 58 GTAGGCGCGGGTGAGGTCATCGCACAGTT 41.8 29 dsrA44 dsrA ACGCGCATGGTATGAAAGTGCGCGACCTT 41.3 29 dsrA45 dsrA GCATGTTCGTCAAGCCGGATCCGTGACGTT 42.2 30 dsrA46 AM179473 dsrA 52 CGTGATGCGATCGCACATCTCGAGCGTGT 41.6 29 dsrA47 dsrA ACTTGTAAGGAAAAGGCGGACGGTGCAGTT 40,0 30 dsrA48a dsrA GTCATTGGGGCACCCCGCGGCCTTGAATT 42.1 29 dsrA49a AM179472 dsrA 40 GAGAAGGTGGGCTCCAGTTGATCCGTTCGC 41.5 30 dsrA50a dsrA TGGTGTGAAAGTGGGCGACCTCGGGGAATT 41.6 30 dsrA51a dsrA GTACTGCATCGTCAGATCGTAGGTCAGGTTGA 41.2 32 dsrA52a AM179464 dsrA 41 GGAGGTGTAGAACCAGGCGCTGGGCTGGT 41.9 29 dsrA53a dsrA CTTCACCTGGGCCTGGTCGATCTGGATGTT 40.4 30 dsrA54 dsrA AGTCGGAACGCGCGATCGATGCGACGC 41.6 27 AY167478, AM179461, dsrA55 dsrA 25 OTU 6 CTGGTATTCCTGCGTCAGGTCGTAAATGATGT 40.4 32 AM179470 dsrA56 dsrA TCGCAACGGGACATTCCCAGGCAGCCCT 41.4 28 dsrA57 dsrA AGAAAACAGGCTCGAGCTGGTCGGTAGTCG 40.9 30 dsrA58 AY167465, AM179459 dsrA 23 OTU 6 GGTCCTCAAGTTGGATCCCGAGCCGCCGA 42.4 29 dsrA59 dsrA GCGCAAGACTTTCGTGTCGTAGTACTTGCT 39.7 30 dsrA60 dsrA CCACAGATCGCACAGACCGCGGATGACTTT 41.6 30 dsrA61 AM179462 dsrA 24 TCCCAGGCAACTTTCCGGAGTCCTGAGGTT 41,0 30 dsrA62 dsrA GCGTCAGATCATAGATGATGTCCTGCGTGTT 39.9 31 dsrA63 AY167469, AM179466, dsrA TTCTCCGGGCAGTCCGAGTACCGGCCGA 41.9 28 31 OTU 3 dsrA64 AY167481 dsrA CAGTCATTGGGACAGGCCGCGACCTTGAT 40.4 29 dsrB65 dsrB CGCGCACCGTATAGAGCGCTTCTCCCGT 41.1 28 dsrB66 AY167469, AY167481 dsrB 32 OTU 3 GATTTCACAGAGATTCGGGATGCGCGTGTT 39.8 30 dsrB67 dsrB GATATCGCATATCTCCCGGATGTGGTCCGTT 40.2 31 dsrA68 dsrA AAGCGGCGTAGGCCGCCACTTCCTTATGG 41.7 29 dsrA69 AM179456 dsrA 33 OTU 3 GATGCAGGCCCACTCACATCGTCCCGGT 40.4 28 dsrA70 dsrA ATAGACCATGCACCTCGTCGGGCAGAGGTT 41.2 30 AY167471, AY167477, dsrA71 dsrA 2 OTU 4 CTTGTACGGGAAAGACGGCCGGTGCATATC 40.7 30 AM179450 AY167471, AY167477, dsrA72 dsrA 3 OTU 4 ATGGTGTCGTAGCAGGCGTATTCGCAGCG 41.2 29 AM179498 AY167477, AM179498, dsrA73 dsrA 4 OTU 4 CTTCCAGACGCCGATGACGCTCATATCGGC 42,0 30 AM179490 AY167477, AM179498, dsrA74 dsrA 5 OTU 4 TCCGGCAGGTCGGAGTAACGGCCGATGAT 42,0 29 AM179465 AY167477, AM179498, dsrA75 dsrA 4 OTU 4 CCCGCAGGGCCTTGGTGGTGTAGAACCAG 41.4 29 AM179490 dsrA76 dsrA GAAGCAGGGCTCCAACTGATCGGTGGTGGT 41.9 30 AM179465, AM179450 6 OTU 4 dsrA77 dsrA GACGGCCGGTGCATATCGAACTGGAATTCC 40.9 30 AY167477, AM179490, dsrA78 dsrA 7 OTU 4 TGGAGCCGTGCTCGTCCCAGATGGCGC 41.3 27 AM179465

70 Sulfate reducers in peatlands

dsrA79 AM179490, AM179465 dsrA 8 OTU 4 CCGGTGCATATCGAACTGGAATTCCTGGGT 39.7 30 dsrB80 dsrB CGAGCTTTTTGCATTCGGCCTTCACGCCG 41.6 29 AY167477, AY167471 9 OTU 4 dsrB81 dsrB TCCACTTGCCGTAGTTCTCTTTGATCACCGG 40.8 31 dsrA82 AM179502, AM179509, dsrA GCGGTGTAATACCAGCCGGAGGGCTGGTT 41.2 29 AM179500, AM179503, 10 dsrA83 AM179499, AM179463 dsrA CATGGTGTGGAAATGTTCCAGACCGGGGAA 39.7 30 AM179502, AM179509, dsrA84 AM179500, AM179503, dsrA 11 GTGGAAATGTTCCAGACCGGGGAACTCATCC 40.8 31 AM179499 dsrA85a AM179463 dsrA 12 GTGGAAATGTTCCAGACCGGGGAAATCATCC 39.9 31 AY167472, AM179474, dsrA86 dsrA 13 OTU11 GGTGCCGATGATGGACATGTCGGCGCGG 42,0 28 AM179460 dsrA87a dsrA CCTTTGCCGTTGTTTCCGCAAAGATGGCTT 40.4 30 AM179474, AM179460 14 OTU11 dsrA88a dsrA GTTGGCGCATCCTGAAACCTTGATCTTGAAC 39.9 31 dsrA89 dsrA CCTTTGCAGTTGTTTCCGCGAAGATGGCCT 41,0 30 AY167472 15 OTU11 dsrA90 dsrA GTTGGCGCAACCGGCAACCTTGATCTTGAA 41.3 30 dsrA91 dsrA TGCCTCGGTCAGTGCATCGAAGCACGGC 41.5 28 AM179506, AM179492, dsrA92 dsrA 39 TGACGCGCACGGTGTGGAAGGCGGTGA 41.6 27 AM179504 dsrA93 dsrA CTTGTACGGCCAGCGGGGTCGATGCATCT 41.6 29 dsrA94a dsrA ATGACATCGCCGGTCGCGCCGTGAAAGTT 42.4 29 dsrA95a dsrA TCATCCTGATACGTCATCGTAAGGTCGTGTATC 40.8 33 AM179475, AM179451 38 dsrA96a dsrA GATGCGCATGGTGTGAAATGCGGTCACGTT 41.8 30 dsrA97a dsrA GATACGTCATCGTAAGGTCGTGTATCAGCGC 40.5 31 AM179506, AM179492, dsrA98 dsrA 39 GATAGGTCATGGTAAGGTCGTGTATCAGCGC 39.8 31 AM179504 dsrA99a dsrA TTAGCTCGTCGAAGCGGGGTTGCAGGCTTT 42.4 30 dsrA100a AM179481 dsrA 36 TGGTATGGAACATCTCAACGGCGGGGAATTTA 41.6 32 dsrA101a dsrA GTGCCGGGCCAACACAGCAACTGATGGTT 41.2 29 dsrA102 dsrA TTCCCAGACATCACAGAGCTTTCTCAAGGTTTT 40.6 33 dsrA103 AM179489 dsrA 37 AAGATTGGCGGTGTTCGTGCCCAAAAGAATG 41,0 31 dsrA104 dsrA ACCTCCGAGATCGAACCCTATCTCTGTCAGTG 41.2 32 dsrA105a dsrA CCAGATATCGCAGAGTCGTCGAAGGTATTTTGT 41.2 33 dsrA106a AM179487 dsrA 35 TCATCGAAGCATGGTTGCAGCTCGTTGGTT 40.8 30 dsrA107a dsrA CACCTGAACCGCCTAAGTCGTAGCCTGCTT 41,0 30 dsrA108 dsrA CCGCATCCTTATTGATACGGATGTCATCTTTCC 40.6 33 dsrA109 AM179488, AM179452 dsrA 17 CAATATCACAGAAACCTCTCAGCCAATCAGTTGT 40.9 34 dsrA110 dsrA CAGGTCCATGGTATCGTAACATGCGAACTCG 40.3 31 dsrA111a AM179485 dsrA 18 CACATGCCCATGGTATCGTAACATGCGAACTC 41.6 32 dsrA112 AM179469 dsrA 19 AACACATGTCCATGGTATCGTAACATACGAACTC 41,0 34 dsrA113 dsrA TACCAATACAATCAGAAGGTGTACGCAGGTTTC 40.7 33 AM179485, AM179469 20 dsrA114 dsrA TGAACGTGCGATACTTGCAGTACAGCCATTC 40.5 31 dsrA115 dsrA CCCTTCTCTCCCACAAGTCACAAATCTTTTCCA 40.9 33 dsrA116 dsrA TATCGTGGGTCAGTTCCCAAAAAACCTCTTCC 40.7 32 AY167464, AY167482 30 OTU 5 dsrB117 dsrB GGCTGCTTGCGGCTCAGGAGATCCTGTTTT 41.5 30 dsrB118 dsrB AATGACCTCGGCGTCGATCATGGGCGGTTT 42.4 30 dsrA119 dsrA CCCCAAACAGCACTCGGGAGTTCTAAGGTTA 40.6 31 dsrB120 AY167475 dsrB 29 OTU 9 GTTTGATCCGCCCGGGAACTTACGGCTTTT 40.7 30 dsrB121 dsrB GCAACCACCTACGCGCACGGAGAAAACCTT 42.1 30 dsrA122 dsrA CAGGTTCGACCCTGAACCGCCCAAGTCCTG 42.4 30 AM179486 28 dsrA123 dsrA TGGTCCGTATTAGTCCCCAGGAAGACGATGTC 41.5 32

71 Part I

dsrA124 dsrA TTGAGGCTGGTCGCAATACCTGCCGATGGT 41.9 30 dsrA125 dsrA CATCAAAGCAGGGCTGAAGATTCTCCGTCTTG 41.1 32 dsrA126 AM179484 dsrA 34 CCCCTAAATCAAAACCCGCATCCGAAAGGTC 40.4 31 dsrA127b dsrA AGCAGATATCCAGGCTGTGGACGCAGGCC 41.1 29 dsrA128 dsrA GCCAGAAGACATAGGGATTGGAACGAGGTTCT 40.8 32 Schlöppnerbrunnen fen soil dsrA129 enrichment clones 2110, 31 dsrA 26 CCAGAGATTCATCAGGTCGCGAAGATTCTTTG 40,0 32 10, 311c dsrB130 dsrB TTTCCATCAATTTCCACCTTGGCGGGCTTTAC 41.2 32 dsrB131a dsrB TCGTTGTCCCCCATGAATTCAACGTTGTTTCT 40.9 32 a Schlöppnerbrunnen fen soil dsrA132 c dsrA 27 GCTTCTGGAAAGAGAGGCGCTTCATGGTTTC 40.6 31 enrichment clone 115RV dsrA133a dsrA CCAGTCCCAGATCTTGCCGACAACGTCTTT 40,0 30 dsrA134 dsrA AGAGGTCGGTGAAGATGTCCTCAAGGTGTTCG 42.1 32 AY167476, AM179476, dsrA135 dsrA 16 OTU 8 GTCGCAGAGATAACGGAGAGAAGACGTGTTGT 41.5 32 AM179494 dsrA136 dsrA GGCATATCGCCATTGTATCCATGCAGGCGT 40.7 30 dsrA137 AM179477, AM179455 dsrA 45 ACCTGAATGTCGTCTTTCCAGGTGCCGAT 38.5 29 dsrA138 dsrA TCAGGTCGTAGCAGAGCTTCATGGTGTCGTA 41.7 31 dsrA139 dsrA CATTCGCGATTCTCGATGGCCAGCTTCTTG 40.3 30 AM179455 46 dsrA140 dsrA CCAGATCGAAACCGGCGTGTGCGAGTTCC 41.6 29 dsrA141 dsrA GCATATTGGTCAGGCCAGACCCGTGCTTCT 41 30 dsrA142 dsrA TCGCGGTTCTCAATGGCGAGCTTCTTGCC 41.5 29 dsrA143 AM179477 dsrA 47 ACTTGTAGGCAAACGCGGGACGATGCAGTT 41.6 30 dsrA144 dsrA CACAGAAGGTGCGCAGGGCATCGCTGGT 41.3 28 AY167465, AM179459, dsrA145 dsrA 21 OTU 6 CGCCAGGTCCCGATAATCGAGCAGTCGGA 41.4 29 AY167478 dsrA146 AM179461, AM179470 dsrA 22 CGCCAGGTCCCGATAATCGAACAGTCGGA 40.3 29 a Perfectlymatched reference target was not available for testing. b Probe was falsenegative during testing and was excluded from further analysis. c unpublished dsrAB clones from a Schlöppnerbrunnen I fen soil enrichment (1) d Affiliation of probetarget group to previously defined Schlöppnerbrunnen dsrABOTU (6)

Table S5. dsrBtargeted primers used for DGGE analysis.

Number of non G+C In silico In silico Primer Sequence (5’3’) degenerated primer PrimerMix (%) coverage (%)a coverage (%)a variants

DSR1728Fa cay acc cag ggy tgg 6073 4 67.0 DSRp1728FmixA 67,0 DSR1728Fb cay acc cag ggr tgg 6073 4 DSR1728FcI cac acb caa ggd tgg 5367 9 6.0 DSRp1728FmixB 6,0 DSR1728FcII cay acb caa ggc tgg 5367 6 DSR1728Fd cat acd cag ggh tgg 5367 4.4 9 DSRp1728FmixC 4,4 DSR1728Ff cac acd cag ggr tgg 6073 6 11.4 DSRp1728FmixD 11,4 DSR1728Fe cac acd cag ggy tgg 6073 6 DSR1728Fg cat acc cag ggn tay 4760 8 5,3 6.3 DSRp1728FmixE DSR1728Fh cat acw cag ggh tat 4047 6 1,0 aCoverage was determined using a customized dsrAB database including 1641 dsrB sequences with sequence information at the primer binding site.

72 Sulfate reducers in peatlands

Table S6. Specificity of dsrABFGA probes as tested by individual hybridization with 36 reference clones. Grey colored boxes indicated hybridization events with normalized signalto background ratios (nSBR) below 0.04 and were considered negative. Yellow boxes indicate hybridization signals between 0.04 and 0.39. Light green boxes specify hybridizations with nSBR between 0.4 and 0.79. Dark green depicts hybridizations with nSBR ≥0.8. Hybridizations between reference clones and perfectlymatching probes are framed in black. False positive hybridizations are thus indicated by colored boxes without black frames, false negative hybridizations by grey boxes with black frames (n=2, probes are additionally indicated in bold).

3_1_1 3_1_1 AY167465 AY167478 AY167475 AY167482 AY167477 AY167466 AY167472 AY167476 AY167469 AY167467 AY167483 AY167474 AM179462 AM179461 AM179486 AM179488 AM179469 AM179490 AM179465 AM179502 AM179456 AM179484 AM179506 AM179489 AM179477 AM179455 AM179495 AM179483 AM179496 AM179510 AM179480 AM179453 AM179473 AM179478 AM179501

dsrA1 0,01 0,03 0,00 0,02 0,01 0,01 0,00 0,01 0,03 0,02 0,01 0,02 0,02 0,25 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,01 0,00 0,02 0,02 0,00 0,03 0,02 0,01 0,00 0,02 0,01 0,02

dsrA2 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,01 0,59 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA3 0,01 0,02 0,01 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,02 0,28 0,60 0,02 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,23 0,01 0,01 0,00 0,01 0,01 0,00 0,02 0,02 0,00 0,00 0,02 0,00 0,01

dsrB4 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,07 0,13 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA5 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,04 0,02 0,36 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,01 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,02

dsrB6 0,01 0,02 0,01 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,24 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,01 0,01

dsrA7 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,01 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,03 0,43 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA8 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,01 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,16 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrB9 0,02 0,02 0,00 0,02 0,01 0,01 0,00 0,01 0,02 0,02 0,01 0,02 0,02 0,00 0,02 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,13 0,00 0,02 0,01 0,00 0,02 0,02 0,00 0,00 0,03 0,00 0,02

dsrB10 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,31 0,27 0,26 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA11 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,07 0,00 0,02 0,01 0,00 0,02 0,01 0,00 0,00 0,03 0,00 0,01

dsrB12 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,05 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA13 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA14 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA15 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA16 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 1,07 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA17 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 1,80 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA18 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,33 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA19 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,20 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrB20 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,36 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA21 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,31 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA22 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,26 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA23 0,01 0,02 0,00 0,02 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,08 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA24 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,17 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA25 0,01 0,02 0,00 0,02 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,11 0,00 0,02 0,00 0,01

dsrA26 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,20 0,00 0,02 0,00 0,01

dsrA27 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,22 0,00 0,02 0,00 0,01

dsrA28 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,06 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA29 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,17 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA30 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,31 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA31 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,09 0,48 0,00 0,00 0,02 0,00 0,01

dsrA32 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,04 0,69 0,00 0,00 0,02 0,00 0,01

dsrA33 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,05 0,01 0,00 0,00 0,02 0,00 0,01

dsrA34 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,09 0,03 0,00 0,00 0,02 0,00 0,01

dsrA35 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 1,15 0,00 0,00 0,02 0,00 0,02

dsrA36 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,81 0,00 0,00 0,02 0,00 0,01

dsrA37 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 1,34

dsrA38 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,40

dsrA39 0,01 0,02 0,00 0,02 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,05 0,14 0,05

dsrB40 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,02 0,01

dsrA41 0,01 0,02 0,00 0,02 0,01 0,01 0,00 0,01 0,02 0,02 0,01 0,02 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,01 0,00 0,02 0,01 0,00 0,02 0,02 0,00 0,00 0,02 0,39 0,01

dsrA42 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 2,20 0,00 0,01

dsrA43 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,01 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 2,15 0,00 0,01

dsrA44 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,09 0,00 0,01

dsrA45 0,01 0,02 0,00 0,01 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,15 0,03 0,00 0,01

dsrA46 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,07 0,03 0,00 0,01

dsrA47 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,05 0,02 0,00 0,01

dsrA48 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA49 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,01 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA50 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA51 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA52 0,01 0,02 0,00 0,01 0,00 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA53 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,01 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA146 0,05 0,02 0,03 0,28 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA54 0,01 0,02 0,20 0,44 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA55 0,01 0,02 0,17 0,70 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA56 0,01 0,02 0,30 1,01 0,01 0,01 0,00 0,01 0,01 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA145 0,35 0,20 0,09 0,10 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA57 0,38 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,01 0,02 0,00 0,01

dsrA58 0,55 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,01 0,00 0,02 0,00 0,01

dsrA59 0,19 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA60 0,01 0,25 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA61 0,01 1,46 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA62 0,01 0,85 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

73 Part I

dsrA63 0,02 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,01 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,30 0,48 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA64 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,31 0,35 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrB65 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,23 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrB66 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,01 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,17 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrB67 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,41 0,95 0,00 0,02 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,02

dsrA68 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,07 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA69 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 1,12 0,00 0,03 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA70 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,15 0,74 0,00 0,02 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrB80 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,01 0,01 0,19 0,16 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrB81 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,41 0,23 0,02 0,01 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,02 0,00 0,00 0,02 0,00 0,01

dsrA71 0,01 0,02 0,00 0,01 0,00 0,01 0,00 0,01 0,02 0,01 0,49 0,02 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA72 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,37 0,04 0,04 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,02 0,00 0,00 0,02 0,00 0,01

dsrA73 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,01 0,01 0,54 0,36 0,04 0,19 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA75 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,01 0,01 0,55 1,34 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA78 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 1,04 1,02 0,73 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA74 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,01 0,01 0,32 0,08 0,14 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA79 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,41 0,36 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA76 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,02 0,03 0,60 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA77 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,63 0,19 0,55 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA82 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,17 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA83 0,01 0,02 0,00 0,01 0,00 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 1,04 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA84 0,01 0,02 0,00 0,01 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,01 0,02 0,00 1,99 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA85 0,01 0,02 0,00 0,02 0,01 0,01 0,00 0,01 0,01 0,01 0,01 0,01 0,02 0,00 0,51 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01 dsrA87 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA88 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA86 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,47 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,03 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01 dsrA89 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,01 0,01 0,01 0,01 0,02 0,00 0,01 0,09 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA90 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,15 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA91 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,55 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01 dsrA92 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,83 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA93 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 1,57 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA98 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,01 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,48 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01 dsrA94 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA95 0,01 0,02 0,00 0,02 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA96 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01 dsrA97 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,01 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA99 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,01 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA100 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,01 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA101 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA102 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,01 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,13 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA103 0,01 0,02 0,00 0,02 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,74 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA104 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,65 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA105 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA106 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,01 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA107 0,01 0,03 0,00 0,02 0,01 0,01 0,00 0,01 0,03 0,02 0,01 0,02 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,01 0,00 0,02 0,02 0,00 0,03 0,02 0,01 0,00 0,03 0,00 0,02

dsrA108 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,14 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA109 0,01 0,04 0,00 0,03 0,01 0,02 0,00 0,01 0,08 0,03 0,01 0,02 0,03 0,01 0,02 0,01 0,01 0,01 0,01 0,00 0,02 0,02 0,01 0,03 0,01 0,00 0,02 0,03 0,01 0,03 0,03 0,01 0,01 0,05 0,01 0,02

dsrA110 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,17 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA111 0,01 0,02 0,00 0,02 0,01 0,01 0,00 0,01 0,02 0,02 0,01 0,02 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,01 0,00 0,01 0,01 0,00 0,02 0,02 0,01 0,00 0,03 0,00 0,02

dsrA112 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,12 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA113 0,01 0,02 0,00 0,02 0,01 0,01 0,00 0,01 0,02 0,04 0,01 0,02 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,01 0,00 0,01 0,01 0,00 0,02 0,02 0,01 0,00 0,03 0,00 0,02

dsrA114 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,17 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA115 0,01 0,03 0,01 0,05 0,01 0,02 0,22 0,01 0,03 0,02 0,01 0,03 0,04 0,01 0,02 0,01 0,01 0,01 0,01 0,00 0,02 0,02 0,01 0,03 0,01 0,00 0,02 0,02 0,00 0,03 0,03 0,01 0,00 0,05 0,00 0,03

dsrA116 0,01 0,02 0,00 0,01 0,01 0,01 0,18 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrB117 0,01 0,03 0,01 0,03 0,01 0,02 0,17 0,01 0,04 0,02 0,01 0,03 0,04 0,01 0,02 0,01 0,01 0,01 0,01 0,00 0,02 0,02 0,01 0,03 0,01 0,00 0,02 0,02 0,01 0,03 0,03 0,01 0,01 0,06 0,01 0,03

dsrB118 0,01 0,02 0,00 0,02 0,01 0,01 0,41 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,02 0,00 0,00 0,02 0,00 0,01

dsrA119 0,01 0,02 0,00 0,01 0,55 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrB120 0,01 0,02 0,00 0,01 0,61 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrB121 0,01 0,02 0,00 0,01 0,40 0,01 0,00 0,01 0,01 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA122 0,01 0,02 0,00 0,01 0,01 0,20 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA123 0,01 0,02 0,00 0,02 0,01 0,13 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,02 0,00 0,00 0,02 0,00 0,01

dsrA124 0,01 0,02 0,00 0,01 0,01 0,05 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA125 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,06 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA126 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,06 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA127 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA128 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,25 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA129 0,01 0,02 0,00 0,02 0,01 0,01 0,00 0,36 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrB130 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,80 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrB131 0,01 0,02 0,00 0,02 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA132 0,01 0,02 0,00 0,01 0,00 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,01 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA133 0,01 0,02 0,00 0,02 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA134 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,30 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA135 0,01 0,03 0,00 0,02 0,01 0,01 0,00 0,01 0,03 0,02 0,01 0,02 0,02 0,01 0,01 0,01 0,10 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,01 0,00 0,01 0,02 0,00 0,02 0,02 0,01 0,00 0,03 0,00 0,02

dsrA136 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,14 0,00 0,01 0,00 0,01 0,01 0,01 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA142 0,01 0,02 0,00 0,01 0,00 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,42 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA143 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,15 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA144 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,60 0,02 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA137 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,20 0,28 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,03 0,00 0,01

dsrA138 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,44 0,67 0,00 0,00 0,06 0,01 0,00 0,02 0,01 0,01 0,00 0,03 0,00 0,01

dsrA139 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,49 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,03 0,00 0,01

dsrA140 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,01 0,01 0,01 0,01 0,02 0,00 0,01 0,01 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,32 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,02 0,00 0,01

dsrA141 0,01 0,02 0,00 0,01 0,01 0,01 0,00 0,01 0,02 0,01 0,01 0,01 0,02 0,00 0,01 0,00 0,00 0,00 0,01 0,00 0,01 0,01 0,01 0,95 0,00 0,00 0,01 0,01 0,00 0,02 0,01 0,00 0,00 0,03 0,00 0,01

74 Sulfate reducers in peatlands

Table S7. Classification of dsrBDGGE sequences in OTUs (based on an amino acid identity threshold of 90%) and corresponding dsrABOTUs from Schlöppnerbrunnen (6) and Rasner Möser wetland sites.

Corresponding OTU dsrB DGGE sequences Schlöppnerbrunnen (Sb) and Rasner Möser (Rm) OTUs 1 DG83 2 DG154 3 DG151 4 DG13 5 DG15, DG16, DG26 6 DG44, DG45, DG46, DG50 7 DG17 8 DG129 9 DG155 10 DG144 11 DG158, DG159 12 DG33 13 DG146 14 DG64, DG65, DG66, DG87 15 DG116 16 DG67 Rm OTU 6 DG112, DG113, DG152, DG153, DG156, DG74, 17 Sb OTU 1 DG79, DG80, DG81 18 DG58 Sb OTU 2 19 DG55, DG101 Sb OTU 3 Rm OTU 1 20 DG91, DG92, DG131 Sb OTU 4 DG35, DG37, DG40, DG51, DG52, DG117, DG123, 21 Sb OTU 6 DG141, DG140 DG18, DG19, DG20, DG21, DG22, DG27, DG31, 22 Sb OTU 11 Rm OTU 11 DG124, DG135, DG82

75 Part I

References

1. Bischof, K. 2003. Vorkommen, Funktion und molekulare Charakterisierung von kultivierten Sulfat reduzierenden Prokaryonten in einem Methan emittierenden sauren Moor. Diploma Thesis. University of Bayreuth, Bayreuth. 2. Haferkamp, I., S. SchmitzEsser, N. Linka, C. Urbany, A. Collingro, M. Wagner, M. Horn, and H. E. Neuhaus. 2004. A candidate NAD+ transporter in an intracellular bacterial symbiont related to Chlamydiae. Nature 432:6225. 3. Haferkamp, I., S. SchmitzEsser, M. Wagner, N. Neigel, M. Horn, and H. E. Neuhaus. 2006. Tapping the nucleotide pool of the host: novel nucleotide carrier of Protochlamydia amoebophila. Mol Microbiol 60:153445. 4. Juretschko, S., G. Timmermann, M. Schmid, K. H. Schleifer, A. PommereningRoser, H. P. Koops, and M. Wagner. 1998. Combined molecular and conventional analyses of nitrifying bacterium diversity in activated sludge: Nitrosococcus mobilis and Nitrospira like bacteria as dominant populations. Appl. Environ. Microbiol. 64:30423051. 5. Kibbe, W. A. 2007. OligoCalc: an online oligonucleotide properties calculator. Nucleic Acids Res 35:W436. 6. Loy, A., K. Küsel, A. Lehner, H. L. Drake, and M. Wagner. 2004. Microarray and functional gene analyses of sulfatereducing prokaryotes in lowsulfate, acidic fens reveal cooccurrence of recognized genera and novel lineages. Appl Environ Microbiol 70:69987009. 7. Schmalenberger, A., H. L. Drake, and K. Küsel. 2007. High unique diversity of sulfatereducing prokaryotes characterized in a depth gradient in an acidic fen. Environ Microbiol 9:131728. 8. SchmitzEsser, S., N. Linka, A. Collingro, C. L. Beier, H. E. Neuhaus, M. Wagner, and M. Horn. 2004. ATP/ADP translocases: a common feature of obligate intracellular amoebal symbionts related to Chlamydiae and Rickettsiae. J Bacteriol 186:68391. 9. Wagner, M., A. J. Roger, J. L. Flax, G. A. Brusseau, and D. A. Stahl. 1998. Phylogeny of dissimilatory sulfite reductases supports an early origin of sulfate respiration. J. Bacteriol. 180:29752982. 10. Zverlov, V., M. Klein, S. Lucker, M. W. Friedrich, J. Kellermann, D. A. Stahl, A. Loy, and M. Wagner. 2005. Lateral gene transfer of dissimilatory (bi)sulfite reductase revisited. J Bacteriol 187:22038.

76

Part II

Hybridization regimes in DNA microarray experiments

DNA microarrays

Introduction

1. DNA Microarrays as high throughput method for community structure analysis

With the ascent of 16S rRNA genebased diagnostic DNA microarrays a powerful method was available to rapidly screen numerous samples. Initially, this tool was used to analyze gene expression patterns (Schena et al. 1995). With the progress in development of the techniques several other applications came up and thus, DNA microarrays are now widely used to profile microbial community structures (Loy et al. 2002; Peplies et al. 2004; Lehner et al. 2005; Loy et al. 2005; Huyghe et al. 2008), to detect pathogens (Wang et al. 2007; WiesingerMayr et al. 2007), or to study bacterial community dynamics and seasonal variations (Brodie et al. 2006; Garrido et al. 2008). In addition, microarrays opened the door to highly parallel structure function analyses of microbial communities with the development of the isotopearray approach (Adamczyk et al. 2003; Wagner et al. 2006). DNA microarrays are based on the parallel detection of tens to thousands different sequence types via multiple hybridization reactions of target sequences with specific probes, which are immobilized on a solid substrate. The DNA microarray technique involves different steps including probe design, microarray fabrication and processing, target preparation, hybridization, and data analysis. Two types of phylochips can be distinguished based on the total number of probes. The high capacity of the microarray format is best exploited with highdensity phylochips that carry several thousands of oligonucleotide probes (DeSantis et al. 2007). In contrast, only up to a few hundred oligonucleotides are immobilized on lowdensity phylochips. These custommade microarrays usually target microorganisms that are defined by their phylogeny or (Loy et al. 2002; Lehner et al. 2005), the environment they live in (Neufeld et al. 2006) or their physiological capabilities. The latter one include also the socalled functional gene arrays (FGAs), which consist of DNA probes targeting genes encoding key enzymes of specific functional or metabolic pathways of microorganisms (Wu et al. 2001) (Bodrossy et al. 2003; Rhee et al. 2004; He et al. 2007; Jaing et al. 2008; Nyyssonen et al. 2009; Waldron et al. 2009).

79 Part II

2. Challenges in DNA microarray experiments

Although largescale comparison of DNA microarray data confirmed an overall high reproducible and reliable data generation by this tool (Shi et al. 2006), substantial technical issues and complex physicochemical processes are associated with the use of this technology, which can hamper the interpretation of microarray data. On one hand, signal variations can originate in the binding behavior and sequence dependence of molecular probetarget hybridizations (Zhang et al. 2003; Wu et al. 2005), the probe length (Chou et al. 2004) or the influence of target folding on heteroduplex formation (Mir et al. 1999). On the other hand, variations such as dye biases caused by dyedye interactions or dyenucleotide interactions (Dobbin et al. 2005; Jeon et al. 2007), spatial artifacts (Yuan et al. 2006) and several other technical variations deriving from microarray manufacturing, sample processing and hybridization (Novak et al. 2002) are known. Systematic variations of retrieved microarray data seem to be one major limitation for correct data analysis, especially if quantitative conclusions are made (Brody et al. 2002; Tan et al. 2003). Underlying mechanisms and their resolution was addressed in several studies, where thermodynamic and kinetic mechanisms such as diffusion limitations or the effect of reaction rates on hybridization were analyzed in several studies (Held et al. 2003; Pappaert et al. 2003; Gadgil et al. 2004; Dandy et al. 2007; Gong et al. 2008; Ono et al. 2008; Singh et al. 2009).

Basically, microarray hybridizations are solidphase hybridizations which are subjected to thermodynamic and kinetic restrictions. The kinetics of hybridizations is mainly determined by mass transport (diffusion) and reaction limited processes (Levicky et al. 2005). Diffusion describes the movement of particles, in the case of microarrays of nucleic acids, with a certain molecular velocity, which depends on the size of the molecule and on the temperature, along a free path. These properties, which are characteristic for different particles in a defined environment, are summarized as diffusion coefficient. Mainly Fick's laws are used to describe diffusion, which add a concentration gradient as driving force to the diffusion coefficient. Microarray experiments are usually dependent on the diffusional transport of labeled nucleic acid targets to their perfect matching complementary probes and hence, diffusion is a key mechanism governing microarray hybridization processes (Dandy et al. 2007). Besides mass transport effects, microarray kinetics also involves hybridization reaction rate considerations, i.e. how quickly the equilibrium of these reactions is approached. Initially, the target DNA strand diffuses towards the immobilized probe strand. During the subsequent collision, a socalled

80 DNA microarrays nucleation site, involving the formation of the first short stretch of base pairing concerning at least 3 contiguous bases, is formed. In the following phase, either a stable base pair is formed during a rapid ‘zippering’ reaction, or, in the case of a mismatch, the nucleation site loses its stability, and the probe strand detaches and prepares for another attempt, either with the same, or with a different probe strand (Southern et al. 1999; Pappaert et al. 2003). In the majority of such reaction rate analyses, the Langmuir isotherm is used as basis for theoretical model development (Halperin et al. 2006). The Langmuir isotherm, originally developed to express the dependence of adsorbed gas molecules on the pressure of the gas above the surface, describes the equilibrium between molecules in solution and corresponding adsorbed molecules bound to the surface of a solid support. To date, mainly the principal nature of hybridization events in microarray experiments was investigated considering single probe–target or single probe spot–target hybridization behavior. Thus, comprehensive analyses of effectors influencing the hybridization mechanisms in relation to the dimensions of such an assay are still missing.

It was the aim of the second part of this thesis to unravel restrictions of DNA microarray experiments by applying a simplified assay format, which allows the analysis of spatial trends and their effectors in DNA microarray hybridizations.

81 Part II

3. References

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Brodie, E. L., T. Z. Desantis, et al. (2006). "Application of a highdensity oligonucleotide microarray approach to study bacterial population dynamics during uranium reduction and reoxidation." Appl Environ Microbiol 72(9): 628898.

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Dandy, D. S., P. Wu, et al. (2007). "Array feature size influences nucleic acid surface capture in DNA microarrays." Proc Natl Acad Sci U S A 104(20): 82238.

DeSantis, T. Z., E. L. Brodie, et al. (2007). "Highdensity universal 16S rRNA microarray analysis reveals broader diversity than typical clone library when sampling the environment." Microb Ecol 53(3): 37183.

Dobbin, K. K., E. S. Kawasaki, et al. (2005). "Characterizing dye bias in microarray experiments." Bioinformatics 21(10): 24307.

Gadgil, C., A. Yeckel, et al. (2004). "A diffusionreaction model for DNA microarray assays." J Biotechnol 114(12): 3145.

Garrido, P., E. GonzalezToril, et al. (2008). "An oligonucleotide prokaryotic acidophile microarray: its validation and its use to monitor seasonal variations in extreme acidic environments with total environmental RNA." Environ Microbiol 10(4): 83650.

Gong, P. and R. Levicky (2008). "DNA surface hybridization regimes." Proc Natl Acad Sci U S A 105(14): 53016.

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Halperin, A., A. Buhot, et al. (2006). "On the hybridization isotherms of DNA microarrays: the Langmuir model and its extensions." Journal of PhysicsCondensed Matter 18(18): S463S490.

He, Z., T. J. Gentry, et al. (2007). "GeoChip: a comprehensive microarray for investigating biogeochemical, ecological and environmental processes." ISME J 1(1): 6777.

Held, G. A., G. Grinstein, et al. (2003). "Modeling of DNA microarray data by using physical properties of hybridization." Proc Natl Acad Sci U S A 100(13): 757580.

Huyghe, A., P. Francois, et al. (2008). "Novel microarray design strategy to study complex bacterial communities." Appl Environ Microbiol 74(6): 187685.

Jaing, C., S. Gardner, et al. (2008). "A functional gene array for detection of bacterial virulence elements." PLoS ONE 3(5): e2163.

Jeon, H. and S. Choi (2007). "Fluorescence Quenching Causes Systematic Dye Bias in Microarray Experiments Using Cyanine Dye." Genomics and Informatics 5(3): 113117.

Lehner, A., A. Loy, et al. (2005). "Oligonucleotide microarray for identification of Enterococcus species." FEMS Microbiol Lett 246(1): 13342.

Levicky, R. and A. Horgan (2005). "Physicochemical perspectives on DNA microarray and biosensor technologies." Trends Biotechnol 23(3): 1439.

Loy, A., A. Lehner, et al. (2002). "Oligonucleotide microarray for 16S rRNA genebased detection of all recognized lineages of sulfatereducing prokaryotes in the environment." Appl. Environ. Microbiol. 68(10): 50645081.

Loy, A., C. Schulz, et al. (2005). "16S rRNA genebased oligonucleotide microarray for environmental monitoring of the betaproteobacterial order "Rhodocyclales"." Appl Environ Microbiol 71(3): 137386.

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Neufeld, J. D., W. W. Mohn, et al. (2006). "Composition of microbial communities in hexachlorocyclohexane (HCH) contaminated soils from Spain revealed with a habitatspecific microarray." Environ Microbiol 8(1): 12640.

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83 Part II

Nyyssonen, M., A. Kapanen, et al. (2009). "Functional genes reveal the intrinsic PAH biodegradation potential in creosotecontaminated groundwater following in situ biostimulation." Appl Microbiol Biotechnol 84(1): 16982.

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84 DNA microarrays

WiesingerMayr, H., K. Vierlinger, et al. (2007). "Identification of human pathogens isolated from blood using microarray hybridisation and signal pattern recognition." BMC Microbiol 7: 78.

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85

Spatial gradients of target nucleic acids govern DNA microarray hybridization assays.

Doris Steger,1 Roman Stocker,2 Susanne Haider, 1 Michael Wagner1 and Alexander Loy1

1Department of Microbial Ecology, University of Vienna, Althanstr. 14, A-1090 Vienna, Austria. 2Department of Civil and Environmental Engineering, Massachusetts Institute of Technology, 77 Massachusetts Avenue, Cambridge, MA 02139, USA.

Draft not corrected by coauthors

Concept by D.S. and A.L. P. amoebophila array conducted by S.H., D.S. performed all other experiments. R.S. performed diffusion modeling. D.S. wrote the manuscript. Part II

Abstract

Nucleic acid surface hybridization assays are routinely applied in science and clinical diagnosis for high-throughput multiplex analyte detection and quantification. This study reports major and systematically arranged spatial gradients of detected target signal intensities within DNA microarrays. An experimental approach shows the general nature of these variations, which halves the amount of detected target in the array center and thus has a major impact on data interpretation and reliability. Additionally, simulation tests unravel mass transfer limitations as causing mechanism.

88 DNA microarrays

Introduction

DNA microarrays, developed in the midnineties (Schena et al. 1995), represent a widely used tool in clinical diagnostics (Yoo et al. 2009), gene expression profiling (Lazazzera 2005; Shyamsundar et al. 2005), genotyping (Sachse et al. 2008; Dufva et al. 2009), microbial ecology (Wagner et al. 2007) or in the detection of alternative splicing (Kechris et al. 2008). They consist of either oligonucleotide or cDNA based probes immobilized on a solid surface and are applied as highthroughput hybridization formats for the identification and quantification of up to several thousand different nucleic acids targets in parallel. It has been shown that DNA microarrays generate overall robust and reliable data with generally high consistencies in identified genes using different microarray platforms (Shi et al. 2006) and thus nowadays array based diagnostic tests are widely spread onto the medical market. However, several studies unraveled systematic variations in microarray data (Brody et al. 2002; Tan et al. 2003) affecting their reproducibility or quantitative power. Possible explanations of such observations can be found in the highly complex physicochemical processes driving these surface capture assays. DNA microarrays are solutionsolidphase hybridizations and thus, underlying dynamics are more complex than in solutionsolution hybridizations. Probes are commonly immobilized on a planar substrate, such as glass slides or nylon membranes, which can interfere with target access (Levicky et al. 2005). Thereby, thermodynamic and kinetic mechanisms regulate nucleic acid hybridizations (Wetmur 1991; Levicky et al. 2005). In DNA microarray experiments two facets of kinetic processes are believed to play a major role: On one hand mass transfer limitations seem to dominate surface hybridizations and on the other hand influence of reaction rates on hybridization has been reported (Held et al. 2003; Pappaert et al. 2003; Gadgil et al. 2004; Dandy et al. 2007; Gong et al. 2008; Ono et al. 2008; Singh et al. 2009). In detail, reaction and/or diffusion modeling of nucleic acid surface hybridization has unraveled diverse chemicophysical phenomena of hybridization events. Pappaert et al. have shown that after an initial reactionlimited time period, target depletion renders the hybridization process diffusion limited (Pappaert et al. 2003). In another study hybridization of short targets was diffusion limited, but long targets were kinetically limited (Singh et al. 2009). Gong et al. reported recently that at low ionic strengths electrostatic forces are dominant, while at high ionic strength more complex hybridization regimes take place (Gong et al. 2008). Studies analyzing probetarget interactions contribute to a better understanding of processes and effectors in hybridization experiments and thus support technical progress in the field and a more accurate interpretation

89 Part II of retrieved data. Thereby, most work focuses on the principal nature of reaction and does not consider spatial scales of such hybridization assays with the exception of one study investigating the influence of microarray spot sizes. In the respective work the authors demonstrated that hybridization efficiencies depend inversely on feature size and increase from spot center to spot edge (Dandy et al. 2007).

The present work bases upon the observation of strong signal intensity gradients across DNA microarrays, which were positioned nonrandomly with significantly lower signal intensities in the array centers. These findings could be confirmed both experimentally by a simplified microarray approach and by diffusion based modeling unveiling strong positional effects in microarray experiments rendering quantitative array data interpretation problematic.

Materials and Methods

Image analyses of Protochlamydia amoebophila array hybridizations and public DNA microarray data.

For the evaluation of systematic variations in microarray experiments, a data obtained by hybridization of a whole genome array for Protochlamydia amoebophila (data unpublished) as well as publically accessible microarray data were analyzed. The P. amoebomophila array consisted of 5´amino modified oligonucleotide probes with a length of 45 to 55 bases. The 5’ end of each oligonucleotide probe was tailed with an additional 20 dATP spacer. Oligonucleotides had a concentration of 100 M in 50% dimethyl sulfoxide (DMSO) and were printed onto silylated aminealdehyde glass slides (VSS25C, CEL Associates, Houston, Tex.) using a BioRobotics MicroGrid spotter (Genomics Solutions, Ann Arbor, MI, USA) and 16 MicroSpot 2500 split pins (Zinsser Analytic GmbH). The average spot diameter was about 150 m and spot centers were 300 m apart. The array consists of 48 blocks, with each block comprising 12 spots per row and column (144 spots/block). The distance between the different blocks was about 1050 m. Analyzed Protochlamydia amoebophila DNA microarrays were hybridized with 1 2 g fluorescently labeled genomic DNA. Briefly, fragmented genomic DNA was random primed and labeled with Cy3 or Cy5

90 DNA microarrays fluorophores by using the DecaLabel DNA Labeling Kit (Fermentas Inc., Hanover, MD, USA). Unincorporated nucleotides were removed using the QIAquick Nucleotide Removal Kit (Qiagen). Purified labeled genomic DNA was vacuumdried in a Speed Vac (Eppendorf concentrator 5301). Before hybridization, slides were prehybridized by addition of blocking reagent in 5x SSC, 0.1% SDS and 1% bovine serum albumin for at least 2 hours at 42°C. Pre hybridized slides were washed in doubledistilled water and in isopropanol, and dried by centrifugation. After denaturation at 95°C for 10 min, labeled targets were hybridized in 400 l of hybridization buffer (35% formamide, 5x SSC, 0.1% SDS, 0.1% nlauryl sarcosine, 0.1% blocking reagent, 50 g ml1 salmon sperm DNA) using sealed coverslips (HybriWell Sealing System; GRACE Bio Labs) under constant shaking at 400 rpm (ThermoTWISTER Comfort, QUANTIFOIL Instruments GmbH). Following 16 hours of incubation at 42°C, the slides were washed at 42°C applying three serial washes with decreasing stringency and images were recorded scanning the slides with a GenePix Personal 4100A array scanner (Axon Instruments, Molecular Devices Corporation, Sunnyvale, CA, USA). Raw hybridization images were used for analysis. The Stanford Microarray database (SMD) was used to evaluate exemplarily public accessible microarray data as it provides a comprehensive and easytouse database including raw images. Analyzed microarrays can be identified by experiment ID (ExptID) given in the figure legends (Demeter et al. 2007). Signal intensity surface plots of microarray hybridization images were generated using the opensource software ImageJ with the Interactive threedimensional Surface Plot plugin (Abramoff et al. 2004). Microarray data were analyzed using the normalized intensities of the average red signal ("R", Cy5) and the average green signal ("G", Cy3) of all open reading frames. Positive and negative controls, as well as additional control oligonucleotides were excluded from analysis.

Microarray system and hybridization experiments

Microarray manufacture and processing. The oligonucleotide, namely DVHO831 (S*Dv.h.o 0831aA18, GAA CCC AAC GGC CCG ACA), for microarray spotting was published previously (Loy et al. 2002) and synthesized from MWG Biotech (Ebersberg, Germany). The 5’ end of each oligonucleotide probe was tailed with 12 dTTP molecules (Tspacer) and the terminal dTTP was aminated to allow covalent coupling of the oligonucleotides to aldehyde groupcoated VSS25 glass slides (CEL Associates). The concentration of probes before

91 Part II printing was adjusted to 50 pmol l1 in 50% DMSO. Oligonucleotides were printed at constant temperature (20°C) and humidity (at least 50%) using a BioRobotics MicroGrid spotter (Genomics Solutions). The DVHO831 probe was printed 900 times per array with 30 features per row and column, respectively. The detailed dimensions of the array are shown in the supplementary Figure 1. Spotted DNA microarrays were incubated overnight at room temperature in a wet chamber. Slides were processed with sodium borohydride as described previously (Loy et al. 2002).

Target preparation. For target preparation, 16S rRNA gene fragments of Desulfovibrio halophilus (DSM5663) were first PCR amplified by the primer pair 616V and 630R (Juretschko et al. 1998) using a concentration of 50 M of each primer. A T3 promoter site tag (5’AAT TAA CCC TCA CTA AAG GG3’) on the 5′ end of the forward primers was added to allow a T3 RNA polymerase based reverse transcription labeling of the PCR products. Reaction mixtures were prepared using 2 mM of each deoxynucleoside triphosphate, 10x Taq buffer, 2 mM MgCl2 and 2 U of Taq DNA polymerase (Fermentas). Cycling was accomplished by an initial denaturation step at 95°C for 3 min, followed by 30 cycles of denaturation at 95°C for 30 s, annealing at 52°C for 30 s, and elongation at 72°C for 1 min. PCR was completed by a final elongation step at 72°C for 3 min. Prior labeling, PCR products were purified using the QIAquick PCR purification kit (QIAgen, Hilden, Germany). Target labeling was achieved by in vitro transcription as follows: 500 ng PCR product, 4 l 5x T3 RNA polymerase buffer (Fermentas), 2 l DTT (100 mM), 0.5 l RNAsin (40 U/l) (Promega), 1 l each of ATP, CTP, GTP (10 mM), 0.5 l UTP (10 mM), 2 l T3 RNA polymerase (20 U/l) (Fermentas) and 0.75 l Cy3UTP or Cy5UTPs (5 mM) in a total volume of 20 l were incubated at 37 °C for 4 hours. DNA was degraded with 2 U DNase I (Fermentas). Enzymatic digestion was stopped with 2 l of 25 mM EDTA (Fermentas). Following adjustment of reaction volume to 100 l with TE buffer, RNA was precipitated with 10

l of 5M NaCl and 300 l of ethanol(abs). RNA was washed with 500 l of ice cold 70% ethanol and resuspended in 50 l H2Obidest. RNA concentration and dye incorporation rates were measured spectrophotometrically. RNA was fragmented by incubating with 10 mM ZnCl2 and 20 mM Tris/HCl (pH 7.4) at 60 °C for 30 min. Fragmentation was stopped by the addition of 10 mM EDTA pH 8.0. Labeled RNA was aliquoted and stored in the dark at 20°C.

Mircoarray hybridization and washing. Prior to microarray hybridization, fluoresecently labeled 16S rRNA of D. halophilus was added to 0.1% SDS, 0.1% 100x Denhardt’s reagent, 6x SSC and 15% formamide and incubated at 65 °C for 515 min. For hybridization, the

92 DNA microarrays

HybriWellTM Hybridization Sealing System (HBW2222FL, Grace BioLabs) with 0.25 mm deep wells was used. Microarray slides were inserted into a custommade hybridization chamber and were hybridized in a waterbath without shaking over night for 18h at 42°C, with the exception of the time series. Subsequently, microarrays were washed by shaking at room temperature for 5 min in 2x SSC, 0.1% (w/v) SDS; for 5 min in 0.1x SSC and finally for 20 s in icecold MQ. Slides were dried by centrifugation (3 min, 300 g) and stored at room temperature in the dark and scanned the same day.

Scanning and data analysis. Fluorescence images were recorded by scanning the slides at three lines to average and at 10m resolution with a GenePix Personal 4100A array scanner (Axon Instruments, Molecular Devices Corporation, Sunnyvale, CA, USA). Photomultiplier tube gain was first adjusted to record spot signal intensities just below the saturation level. For each experimental series, identical scanning settings were used to enable the comparison of absolute signal intensities between different hybridizations. Scanned images were saved as multilayer tiff images and analyzed with the GenePix Pro 6.0 software (Axon Instruments). Low quality hybridizations were repeated and single bad quality spots were excluded from analysis. Signal intensity surface plots of the microarray results were generated using the tool ImageJ with the Interactive threedimensional Surface Plot plugin (Abramoff et al. 2004). Signal intensities were analyzed according to the defined spot positions as illustrated in supplementary Figure 1. For data analysis, median pixel intensities of probe spots were used. Mean relative signal intensities for each spot position were calculated as ratio of the spot position with highest or lowest absolute signal intensity, respectively. Oneway analysis of variance (ANOVA) was used to compare the mean differences of signal intensities of different spot positions.

Results and Discussion

Spatial gradients in microarray experiments.

Image analysis of hybridization experiments with fluorescently labeled genomic DNA on a whole genome array of Protochlamydia amoebophila pointed to periodic intraarray variations in intensities of fluorescent signals (data unpublished). An obvious increase in detected fluorescence intensities from the array center to the array edge was observed via interpretation

93 Part II of absolute signal intensity as height in a three dimensional surface plot (Figure 1). These systematic differences in signal intensities could be observed both, over the whole width of the microarray (Figure 1A) and within the single subblocks (Figure 1B), respectively.

A B

Figure 1: Image analysis of a hybridizations with fluorescently labeled genomic DNA on a genome array of Parachlamydia pneumophila. Signal intensities are shown as height in a three dimensional surface plot using ImageJ and displayed as (A) lateral view and (B) top view.

Additionally, publically accessible microarray experiments, which were hybridized with fluorescently labeled genomic DNA, exhibited comparable variations in signal intensities with considerably higher signal intensities at the border of the array or the array subblocks than in the array or block centers (supplementary Figure 2) (Demeter et al. 2007). These findings indicate that observed phenomenon concerns not only the P. amoebophila array but could be of common relevance in microarray hybridizations.

In order to verify and characterize the nature of the observed hybridization behavior, a simplified array layout was chosen, which allows a reduction in complexity of putative influencing factors. Therefore, the same oligonucleotide probe was immobilized in 900 spots (30 spots per row and column) to exclude different reaction rates of dissimilar probes as driving force for variability in

94 DNA microarrays signal intensities. The microarray was hybridized with varying concentrations of target RNA for different time periods (18, 65 and 140 hours) without agitation. Figure 2 shows hybridization of Cy3 labeled RNA under targetlimiting conditions (300 ng) for 18 hours, a typical hybridization setup in DNA microarray experiments (Ulger et al. 2003; Bodrossy et al. 2006; Han et al. 2006; Manak et al. 2006). A B

1.2

1.1

1. 0 0.9

0.8

0.7

0.6 relative signal intensity signal relative 0.5

1 2 3 4 5 6 7 8 9 1011121314 15 spot position Figure 2: Analysis of target hybridizations on a simplified assay under target limiting conditions for 18 hours. Fluorescently labeled RNA was hybridized on an array with 900 spotted features consisting of 18mer oligonucleotides complementary to the target. (A) Overlay of 3 replicate hybridizations using Cy3 labeled RNA Mean signal intensities are shown as height in a three dimensional surface plot using ImageJ (B) Microarray hybridization of Cy3 (red) and Cy5 (blue) labeled target RNA. Mean relative signal intensities are given as fraction of the spot position with highest absolute signal intensity and are displayed as function of the respective position. Error bars represent relative standard deviations of the mean signal intensities per spot position (including values of 3 independent experiments).

Comparison of the different spot positions as defined in supplementary Figure 1 revealed a significant decline of detected amount of hybridized target (p < 0.001) from the array edge (spot position 1) to the array center (spot position 15) (Figure 2B). If measured signal intensities were expressed as fraction of the strongest signal, mean signal intensities in the array center almost halved in relation to those at the array edge (Figure 2A). To exclude dye specific effects, equivalent hybridizations were carried out using Cy5 instead of Cy3 labeled RNA and obtained results corresponded to those received in the former experiment (Figure 2B) (Pearson correlation coefficient between Cy3 and Cy5 signals of the different spot positions = 0.99, p < 0.01). Moreover, after comparison of apparent differences in signal intensities within the distinct

95 Part II spot positions, highest values in the array corners were detected (Figure 3). By considering the signal intensities of the diagonal and center line transect separately, significant higher amounts of hybridized target could be observed at the outer spots of the diagonal transect compared to the corresponding features of the center line transect (p < 0.01).

2.4

2.2

2.0

1.8

1.6

1.4 1.2 intensity signal relative 1.0

1 2 3 4 5 6 7 8 9 101112131415

spot position

Figure 3: Comparison of relative signal intensities of diagonal (black) and center line transects (grey) (see supplementary Figure 1) of a hybridization with Cy3 labeled RNA under target limiting conditions for 18 hours. Mean relative signal intensities are given as fraction of the spot position with lowest absolute signal intensity and are displayed as function of the respective position. Error bars represent relative standard deviations of the mean signal intensities per spot position (including values of 3 independent experiments).

It can be assumed, that spots at the array corners have a positional advantage to access the provided target. As they have fewer spots in their direct surroundings, they have less competition for RNA strands in solution. Moreover, physical interference by other adjacent probe spots hindering the free diffusion of the target is reduced. Our results strongly point to mass transfer limitations as key driver in such surface hybridizations as the same probe was spotted at all position and thus different probetarget reaction kinetics can be excluded. In a study of Livshits et al., brighter signals in DNA hybridization assays with oligonucleotides immobilized in a polyacrylamide gel layer have been observed at the edges. Retarded diffusion was shown to direct the kinetics of hybridization on the investigated assay (Livshits et al. 1996). Essentially, the movement of DNA into the center of the assay was shown to be driven by diffusion, whereby repeated duplex formation between probes and free analyte molecules is assumed (Livshits et al. 1996). As a result their diffusion is slowing. In a more recent work,

96 DNA microarrays spatial biases in DNA microarray experiments were described as a major source of inaccuracies in such studies (Koren et al. 2007). Although this study did not provide any explanation for the observed variations, they could show that the described biases might be responsible for more than 15% false data per experiment independent of microarray platforms and experimental procedures.

1.0 1.0

0.9 0.9

0.8 0.8

0.7 0.7

0.6 0.6 relative relative signal intensity relative relative signal intensity 0.5 0.5 1 2 3 4 5 6 7 8 9101112131415 75 150 300 600 1800 4500 spot position concentration (ng)

4500 ng 1800 ng 600 ng spot positions 1 2 3 4 5 6 7 8 300 ng 150 ng 75 ng 9 10 11 12 13 14 15

Figure 4: (A) Hybridization series with different concentrations of Cy3 labeled target RNA (75 – 4,500 ng) for 18 hours. Mean relative signal intensities are given as fraction of the spot position with highest absolute signal intensity and are displayed as function of the respective position. Error bars are omitted for clarity. Relative standard deviations of the mean signal intensities per spot position (including values of three independent experiments) ranged between 0.22–0.08 for 75 ng, 0.21–0.13 for 150 ng, 0.20–0.10 for 300 ng, 0.20–0.11for 600 ng, 0.12–0.7 for 1,800 ng and 0.12–0.7 for 4,500 ng, respectively. (B) Comparison of relative signal intensities for the different spot positions using different target amounts. Mean relative signal intensities (including 3 independent experiments) are given as fraction of the spot position with highest absolute signal intensity and are displayed as concentration.

To further investigate probe–target hybridizations on a surface, a series of experiments using several target concentrations were conducted. A remarkable decrease of spatial gradients at 1800 ng and their almost complete disappearance using 4,500 ng of target rRNA was observed (Figure 4A). Comparison of the different target amounts hybridized to the distinct spot positions mirror their dissimilar hybridization behavior at respective spot locations (Figure 4B). This finding is not surprising, as mass transfer and hybridization rates, both key processes in

97 Part II hybridization experiments, are strongly dependent on the concentrations of analyzed targets (Bhanot et al. 2003; Dandy et al. 2007). Under target limiting conditions, a similar picture is presented by the time series with a decrease in variation of the signal intensities for the different spot positions over time (Figure 5A). A closer look at the single spot positions over different time confirms a heterogeneous hybridization behavior of target RNA to the immobilized probe spots (Figure 5B).

A B

1.0 1.0

0.9 0.9

0.8 0.8

0.7 0.7

0.6 0.6 relative relative signal intensity relative relative signal intensity 0.5 0.5 1 2 3 4 5 6 7 8 9 101112131415 18 65 140 spot position time (h) 18 h 65 h 140 h spot positions 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15

Figure 5: (A) Hybridization series of Cy3 labeled target RNA under target limiting conditions for 18, 65 and 140 hours. Mean relative signal intensities are given as fraction of the spot position with highest absolute signal intensity and are displayed as function of the respective position. Error bars are omitted for clarity. Relative standard deviations of the mean signal intensities per spot position (including values of three independent experiments) ranged between 0.20–0.10 for 18 h, 0.12–0.04 for 65 h and 0.21–0.11 for 140h, respectively. (B) Comparison of relative signal intensities for the different spot positions over time under target limiting conditions (Cy3 labeled RNA). Mean relative signal intensities (including 3 independent experiments) are given as fraction of the spot position with highest absolute signal intensity and are displayed as function of time.

While the inner spot positions 11 to 15 display a constant increase in signal intensities, the slope diminishes for the positions 2 to 10. Finally, the outer spot position 1 demonstrates no obvious differences in the amount of hybridized target over time with a tendency to decreasing signal intensities after 65 hours. These results underpin the assumption of mainly diffusion limiting transport. If the overall hybridization performance on the array would be mostly reaction

98 DNA microarrays rate limited, the same probetarget hybridization behavior would be expected at all spot positions. Thus, simultaneous presence of different reaction states within one array is likely. In the case of spot position 1, the amount of hybridized targets does not change significantly over time. Accordingly, reaction rate decreases suggesting depletion of available free analyte strands in the surroundings of these probes or saturation of the spots at position 1, respectively. In contrast, spot hybridization reactions in the array center seem to be distant from steady state, likely caused by target limitation and continuous new influx of free targets by diffusion.

A ratio B spot signal intensity position (Cy3/Cy5) 120 1 2.5 1.1 2 2.6 100 3 2.7 1.0 4 2.7 80 5 2.8 0.9 6 2.8 60 7 2.8 8 2.8 0.8 40 9 2.9 0.7 10 2.9

normalized Cy5normalized intensity 20 relative signal signal intensity relative 11 2.9 12 2.9 0.6 1 2 3 4 5 6 7 8 9 101112131415 13 2.9 20 40 60 80 100 120 14 2.9 spot position normalized Cy3 intensity 15 2.9

Figure 6: Analysis of simultaneously hybridized Cy3 and Cy5 labeled targets on DNA microarrays. (A) Double hybridization of 900 ng Cy3 (red) and 300 ng Cy5 (blue) labeled target RNA on the simplified assay (see supplementary Figure 1) for 18 hours. Mean relative signal intensities are given as fraction of the spot position with highest absolute signal intensity and are displayed as function of the respective position. Error bars represent relative standard deviations of the mean signal intensities per spot position. Additionally, ratios of Cy3 and Cy5 median pixel intensities per probe spot positions are shown. (B) Linear scatter plot of a double genomic DNA hybridization (ExpID 68809). Each ORF is represented by a point with coordinates consisting of the normalized intensities of the average Cy5 signals and Cy3 signals. The array consisted of 12 subblocks with eight blocks located on the borders of the assay and four in the assay center (compare with supplementary Figure 2A). ORFs located in the outer blocks are displayed in red (n=2882), ORFs of the array center in black (n=959).

Finally, double hybridization of varying Cy3 (900 ng) and Cy5 (300 ng) labeled RNA concentrations also reflects a differentiated hybridization behavior. The continuous decrease of hybridized target amount to complementary probes of the spot positions 1 to 15 was lower for the applied threefold higher concentration of Cy3labeled target strands than for the Cy5 labeled targets (Figure 6A). This heterogeneous hybridization behavior is a crucial factor in

99 Part II microarray data interpretation, as gene expression array experiments usually include different concentrations of targets, ranging from abundant to rare target strands. Highly concentrated targets are closer to steady state than targets that have a low concentration level. Consequently, an asymmetric precision in the identification of up and downregulated genes was anticipated (Bhanot et al. 2003). In detail, lower target concentrations correspond usually to downregulated genes and according to their extended equilibrium times, longer hybridizations would be needed to obtain the same accuracy as for genes that are upregulated (Sartor et al. 2004). Hybridization times needed to reach equilibrium depends on target concentration and thus, underestimation of fold change of differentially expressed genes and miss of low abundant genes is suggested when hybridization reactions are not driven to completion (Wei et al. 2005). Furthermore, comparison of the actual applied target ratio of 3:1 with the experimentally measured Cy3toCy5 ratios, fitted best in the center of the array (2.9:1, spot position 9 to 15), where targetlimiting and nonequilibrium conditions are assumed (Figure 6A). It has been shown that linearity between target concentrations and measured amounts of hybridized targets is sustained only within a narrow concentration range. Thereby very low abundant targets are affected by detection limit issues and high abundant targets are influenced by saturation penalties (Chudin et al. 2002). Most studies emphasize the importance of equilibrium conditions for adequate quantification of target molecules, especially in regard to multiplex hybridization as crosshybridizations are enhanced under nonequilibrium condition (Bhanot et al. 2003). Nevertheless, our results suggest best agreement of actually present and hybridized target concentrations under nonequilibrium conditions. Accordingly, theoretical modeling by Gadgil et al. supports this finding in diffusion driven reactions, since intensities of hybridized targets corresponded best to their actual concentrations, when the reaction was far from equilibrium (Gadgil et al. 2004). In order to verify these findings, public accessible data of a genome microarray hybridized simultaneously with Cy3 (1g) and Cy5 (4 g) labeled genomic DNA of different cholerae strains were analyzed exemplarily using the normalized intensities of the average Cy5 and Cy3 signals of all open reading frames. By comparison of the Cy5 and Cy3 signal intensities of the subblocks positioned on the boundaries of the array against those located in the array center, an uneven distribution of analyzed signal intensities was observed. In detail, spots positioned in the array center possess a narrow range of Cy3toCy5ratios, whereas the outer subblocks exhibited a broader distribution with an apparent overrepresentation of Cy5 labeled strands hybridized on the respective spots (Figure 6B). This analysis underlines the importance of the observed effect in real microarray experiments, such as gene expression analyses, as multiple

100 DNA microarrays targets with different specificities and concentrations appear to be affected by the same hybridization mechanisms than our simplified array format. Although the effect can be influenced by the sequence composition and thus by unequal hybridization behavior of the two hybridized strains, a preferential hybridization driven by higher affinities of the Cy5 labeled V. cholerae strain only to spots of the outer blocks seems to be highly unlikely.

Diffusion drives the hybridization process.

The confirmation of the obtained experimental data was accomplished by simulation of the passive mass transport using a diffusion model. It was presumed that target transport is only driven by Fickian diffusion kinetics. Thereby, the diffusion equation was solved for the simplified array format used for the experiments with the dimensions as illustrated in supplementary Figure 1, with the exception that the chamber height assumed in the numerical simulation was 150 m and 750 m, respectively. In detail, a hybridization chamber of a halfwidth L=10.8 cm and a height H=150 m, containing a square block of 30 x 30 spots (0.85 x 0.85 cm), each spot having 150 m in diameters and a distance of 150 m between them, is assumed. A time dependent simulation with a diffusion coefficient of D=109 m2/s and a diffusion time of 86400 s was applied. The flux of target strands to each spot at each instant in time was calculated and was subsequently integrated up for the total time (24 h) for each spot. Thereby, three different cases were considered. Case A is most similar to the above described situation on the applied simplified array with a large and shallow hybridization chamber (L=10.8 mm, H=150 m), with the exception that the actual chamber height was 250 m. Case B assumes a large and deep hybridization chamber (L=10.8 mm, H=750 m) and is used to study the effect of the chamber’s height. Finally, case C tests the effect of the amount of surface area not covered with spots by assuming a small and shallow chamber (L=4.5 mm, H=150 m).

101 Part II

1.2

1.0

0.8

0.6

0.4

0.2 Flux (relative Flux (relative tospot position 1)

1 2 3 4 5 6 7 8 9 101112131415 spot position

Figure 7: Numerical simulations of target diffusion relative to the spot positions as defined in supplementary Figure 1. The flux of target strands to each spot position was estimated for a hybridization time of 24 h. Three different cases were considered: Case A (red) corresponds to an array with a large and shallow hybridization chamber (L=10.8 mm, H=150 m). Case B (dark blue) assumes a large and deep hybridization chamber (L=10.8 mm, H=750 m) and case C (light blue) assumes a small and shallow chamber (L=4.5 mm, H=150 m).

Resulting profiles displayed as relative flux of target RNA to the respective spot positions (Figure 7) revealed a significant decrease in the amount of RNA transported to the spots at the array center considering case A and B, and thus confirmed the results obtained in the hybridization experiments. Thereby, chamber height influences the amount of target molecules reaching the array center. In detail, the shallower the hybridization chamber is, the more pronounced the spatial variability of hybridized target will be. In fact, when the hybridization chamber is very shallow, most target molecules will hybridize to the outer spots before they can penetrate the block, whereas in deeper ones changes of RNA getting "captured" by the outer spots is lower. Actually, diffusionreaction modeling of DNA hybridizations on microarray showed a linear increase of hybridized targets with chamber height. Furthermore it was implied that the time range for the change from reaction to diffusion limited hybridization is a function of chamber height (Pappaert et al. 2003). Thereby, targets within the first 100 nm of the fluid layer above the probe spots were fully hybridized in 10 ms, whereas the depletion of target strands within 100 m took 1000 s, as the process switched to a diffusion controlled regime. Furthermore, in an experimental study, free targets appeared to be quickly depleted perpendicular to the array. By increasing the chamber height, greater advantage of vertical diffusion is taken, and thus the shift to a lateral diffusion limited hybridization is delayed (Borden et al. 2005).

102 DNA microarrays

Finally, comparison of case A and C uncovered the presence of a free surface area not covered with probe spots within the hybridization chamber as crucial factor in determining the spatial variability (Figure 7). If the chamber included no spotfree surface, no spatial variability was estimated (case C). Accordingly, each spot depletes rapidly all free targets in its neighborhood (over a time scale of approximately H2/D, i.e. 9.3 minutes for a height of 750 m). Thereby, all spots have equal access conditions to free target strands perpendicular to their positions. In contrast, when an uncovered surface area is included in the array, which in our case was 6.38 cm, target strands will diffuse for the most part laterally towards the spotted area, and will therefore hybridize preferentially to the outer spots in the block. The differences of size of the applied hybridization chamber with a halfwidth within cm and height within mscale, respectively, governs observed hybridization regimes.

103 Part II

Conclusions

Spatial gradients present in nucleic acid surface hybridizations were investigated systematically by hybridization experiments and numerical simulations. A simplified microarray approach confirmed the preferential hybridization of analytes at the array edges resulting in 50% lower signal intensities in the array center at the given array dimensions. Observed gradients appeared also in twocolor analyses, usually applied in gene expression analysis. As a consequence, quantitative data interpretation is heavily biased by systematic spatial variations. Theoretical simulations emphasized these results and identified diffusion processes as key driver for the heterogeneous hybridization behavior at different spot positions.

This study reports and elucidates a drastic and systematic spatial effect tampering certainly most nucleic acid surface hybridization experiments. Reduction of the influence of described variations could be approached by the adaptation of already existing normalization methods such as integration of embedded and adequately distributed technical replicates for bias correction (Yuan et al. 2006). Furthermore, reduction of diffusion driven reactions is approached by the application of non static hybridization devices. Most commercial DNA microarray formats, including Affymetrix Gene Chips, utilize only passive mixing by rotation of the entire microarray, and thus still suffer from diffusionlimited hybridization rates. However, possibilities to overcome diffusion limitations are provided by actively mixing hybridization systems. One promising attempt is the use of pressure driven flowthrough systems. The porous substrate was shown to reduce dramatically diffusion limitations on the hybridization rate (Mocanu et al. 2009); nevertheless, further effort will be needed to unravel the complex physicochemical interactions in these relatively novel assays. Finally, a straightforward improvement was highlighted by the simulation experiments. The extension of the height and contemporary reduction of free surface areas within the array would be an easy and effective way to allow faster and equal analyte transfer to all spots. However, larger hybridization solution volumes would be required and therefore the detection of low abundant target might be lowered.

104 DNA microarrays

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Supplementary Figures

hybridization chamber

array L H

30 spots diagonal transect 0.15 mm0.15

0.15 mm 30 spots 8.85 mm center line 15 transect 14 13 12 11 10 9 8 7 6 5 4 3 2 1 8.85 mm

Supplementary Figure 1: Assay layout and dimensions. Hybridization chambers have a half width of L=10.8 cm and a height of H=0.150 cm. Spot positions (1–15) are labeled with white and grey surrounds. Features within one defined position are treated as replicate spots with decreasing numbers of replicates from spot position 1 (n=116) to spot position 15 (n=4). Definitions of diagonal transects (n=4) and center line transects (n=8) are shown as dotted lines. Spots have a diameter of 150 m and have a distance of 150 m to each other.

109 Part II

A B

Supplementary Figure 2: Image analysis of public accessible microarray images. Analyzed microarrays can be identified by the experiment ID (ExptID) in the Stanford Microarray database. Genomic DNA hybridizations on different target genome arrays were chosen for analysis. Signal intensities are shown as height in a three dimensional surface plot using ImageJ. (A) ExpID 68809 (B) ExpID 75165.

110

Part III

Microorganisms in marine sponges

Image from www.ramblincameras.com (© WBM Enterprises)

Introduction

Sponges (Phylum Porifera) are the most primitive of the multi-cellular (Metazoa). They form one of the deepest branches of the metazoans, whose evolutionary roots date back to times over 600 million years ago (Müller et al. 2009). There exist about 15000 known and 8000 validly described species with the vast majority (85%) of the formally described species belonging to the class Demospongiae (Hooper et al. 2002). These sessile, filter-feeding organisms inhabit a wide variety of marine and freshwater systems. Sponges frequently colonize tropical reefs but are also found in temperate and polar regions as well as in freshwater streams and lakes. They represent important members of benthic communities, as they contribute substantially to the present in the benthic habitat such as reefs (Diaz et al. 2001). In this connection, marine sponges play an important functional role in benthic or pelagic processes, as they offer a habitat for various reef species and they are heavily involved in benthic food webs (Gili et al. 1998; Diaz et al. 2001). Two key factors direct research interest towards marine sponges: on one hand marine sponges are recognized for their production of secondary metabolites with a wide range of biological and pharmaceutical activities (Braekman et al. 2004; Laport et al. 2009) and on the other hand many species harbor dense and highly diverse microbial communities (Hentschel et al. 2006; Webster et al. 2009) which have been postulated to be important for the fitness of the and which might contribute to the production of secondary metabolites (Piel 2004; Wang 2006; Muscholl-Silberhorn et al. 2008; Hertiani et al. 2010).

1. Microorganisms associated with marine sponges

To date, several types of marine sponge-microbe associations are known. Microorganisms are used as food source by these filter-feeding organisms, but they comprise also disease- causing parasites or mutualistic symbionts (Wilkinson et al. 1984; Webster et al. 2002; Müller et al. 2004; Thacker 2005). This variety in the nature of interactions is also reflected by the phylogenetic diversity of microorganisms that occur within sponges: all three domains of life, namely Bacteria, Archaea and Eukarya, are now known to inhabit marine sponges. Microorganisms occur both intracellularly in specialized sponge host cells, the bacteriocytes, and extracellularly in the sponge mesohyl (Friedrich et al. 1999). Thereby, intracellular sponge symbionts are less frequently observed then the often dense microbial communities

113 inhabiting the sponge mesohyl, an extracellular matrix which separates the internal acquiferous systems from the outer sponge cells. Twenty-three bacterial phyla, both major archaeal lineages, and various microbial have been described so far in marine sponges (Webster et al. 2001; Taylor et al. 2007; Granados et al. 2008; Webster et al. 2009). Thus, marine sponges contain highly diverse microbial communities with often immense amounts of microorganisms detected within one sponge (up to 109 microbial cells per g) (Hentschel et al. 2006). Since microbes detectable in sponges can either be symbionts or a food source, it is challenging to reliably assign them to one of these two groups.

One approach to distinguish between the different types of sponge associated microorganisms employs the phylogenetic characterization of these associations. For instance, comparative taxonomic analysis of selected marine sponges and their bacterial associates suggested that some sponge symbionts have co-evolved with their hosts, implying a permanent and maybe beneficial role of these bacterial groups (Erpenbeck et al. 2002). Furthermore, numerous sponge associated microorganisms were affiliated to sponge-specific clusters, based on their 16S rRNA gene sequences (Hentschel et al. 2002), suggesting selective occurrence of certain bacterial and archaeal groups within sponge hosts. Per definition, these clusters have to fulfill the following conditions: they have to comprise at least three microbial sequences that (i) have been retrieved from different sponges and/or from distinct geographic locations, (ii) are more closely related to each other than to any other sequence from non-sponge sources, and finally (iii) cluster together independent of the treeing method used (Hentschel et al. 2002). Out of a total number of 190 analyzed 16S rRNA gene sequences, 133 sponge-derived sequences (70%) were originally assigned to 14 monophyletic sponge-specific clusters in 2002. This basis was extended comprehensively in the course of this thesis in 2007 by phylogenetic analyses of about 1700 microbial sponge-derived 16S rRNA gene sequences with 546 16S rRNA gene sequences affiliating to sponge-specific clusters (32%). Recent bacterial 16S rRNA gene tag pyrosequencing of three different marine sponge species and surrounding samples confirmed the validity of 48% out of 33 detected clusters (Webster et al. 2009). Some of the remaining sponge-specific clusters were also found at very low numbers in the surrounding seawater. Consequently, the rare seawater biosphere possibly provides microbes as seed organisms to sponges resulting in selective enrichment of specific microbes from the marine environment. Thus, described host-microbe associations might have evolved much more recently than previously thought (Hentschel et al. 2002; Webster et al. 2009).

114 Beside the possibility of environmental acquisition of microorganisms by the host sponge, evidence of vertical transmitted bacteria via sponge larvae was obtained in several studies, which supports stable associations of microorganisms in at least some sponges. A first insight into vertical transmission in marine sponges was provided by ultrastructural studies. The presence of located throughout the mesohyl of adult Chondrilla australiensis sponges, was additionally shown inside developing eggs and nurse cells in 25% of hosts using transmission electron (Usher et al. 2001). Furthermore, yeast cells were maternally transmitted in Chondrilla species through the oocytes to the fertilized eggs (Maldonado et al. 2005). Following molecular studies revealed that multiple and phylogenetically diverse groups of microorganisms are transferred between sponge generations through their reproductive stages (Enticknap et al. 2006; Schmitt et al. 2007; Sharp et al. 2007; Schmitt et al. 2008; Lee et al. 2009). Combined application of 16S rRNA gene analyses, fluorescence in situ hybridization and electron microscopy resulted in a model for symbiont transmission in marine sponges with high microbial abundances (Schmitt et al. 2008). In this model sponge reproductive stages take up microbes from the adult mesohyl. Nurse cells might be involved essentially in microbial incorporation by promoting selection of these microbial communities. Developing sponge larvae are non-filter feeding stages and might be regarded as a closed system, where no exchange with the environment occurs. Vertical transmission leads consequently to separation of microbial types in different sponges which would result in genetic differentiation and finally in co- speciation. After the metamorphosis of sponge larvae into juveniles, the filter-feeding life phase begins, which induces uptake of microorganisms from the surrounding seawater (Schmitt et al. 2008). Thereby, horizontal transfer of microbes between sponge individuals or environmental acquisition as discussed above appears to be possible.

2. Ecological functions of sponge associated microorganisms

There are a variety of functions that have been attributed to microorganisms living in marine sponges. Host sponges may benefit from microbial metabolite production, sponge stabilization, nutrient acquisition and metabolic waste removal (Wilkinson 1978; Wilkinson 1992; Unson et al. 1994; Hentschel et al. 2001). For instance, symbiont-produced bioactive metabolites may serve to protect the sponge from pathogens, predators and organisms (Hentschel et al. 2001). Furthermore, facultatively anaerobic bacteria are commonly found in sponges (Wilkinson 1978; Santavy et al. 1990) and through their ability

115 to metabolize a wide range of compounds, these organisms may play a role in removing waste products from the sponge during periods of low pumping activity (Wilkinson 1978). A well studied example of sponge-microbe interactions is presented in the beneficial contribution of cyanobacterial symbionts to their host. Many tropical sponges contain substantial populations of these oxygenic photoautotrophs and it was demonstrated that photosynthetic produced glycerol and organic phosphate are provided from cyanobacterial symbionts to their hosts (Wilkinson 1979). In this case, the host sponge may receive up to 50% of its energy requirements and 80% of its carbon budget (Wilkinson 1992; Cheshire et al. 1997). A further advantage for the host may include the provision of nitrogen, a limiting nutrient in many marine systems, by cyanobacteria which are capable of fixing atmospheric nitrogen (Wilkinson et al. 1979).

Further microorganisms involved in the nitrogen cycle have been identified in marine sponges. The nitrogen cycle in sponges comprises four main microbial mediated transformation processes, namely , nitrification, anaerobic ammonia oxidation and denitrification. During nitrogen fixation, carried out by specialized microorganisms like the above mentioned Cyanobacteria, nitrogen is converted to ammonia. Under aerobic conditions, ammonia is transformed to nitrite and subsequently to nitrate by ammonia- and nitrite-oxidizing microorganisms (nitrification). Under anaerobic conditions however, members of the genus Planctomycetales are able to convert nitrite and ammonium directly into N2 in a process called anaerobic ammonia oxidation (Jetten et al. 2001). Finally, nitrate and nitrite can be reduced anaerobically to N2 (or NO and N2O) by diverse heterotrophic bacteria (denitrification). It has been shown, that marine sponges assimilate and release dissolved inorganic nitrogen like ammonia, nitrite and nitrate, as well as dissolved and particulate organic nitrogen (Bayer et al. 2008). In addition, microbial mediated denitrification and anaerobic ammonium oxidation in sponges was quantified by application of incubation experiments with stable isotopes (Hoffmann et al. 2009). Also molecular methods were employed using 16S rRNA genes and functional genes as phylogenetic markers, to uncover the microorganisms involved in these transformations. Environmental genome sequencing of the sponge associated crenarchaeon Cenarchaeum symbiosum reported the presence of an ammonia monooxgenase gene (amoA), a key enzyme in the aerobic ammonia oxidation, in this uncultivable sponge symbiont (Hallam et al. 2006). Additionally, 16S rRNA gene and/or amoA genes of Nitrosospira, Nitrospira and Crenarchaeota, all groups involved in aerobic nitrification, were detected in several marine sponges (Hentschel et al. 2002; Bayer et al. 2008; Hoffmann et al. 2009; Mohamed et al. 2009). Recently, 16S rRNA gene based analyses revealed furthermore first molecular evidence for the presence of anaerobic ammonia oxidizing bacteria in sponges (Hoffmann et

116 al. 2009; Mohamed et al. 2009). Finally, investigation of functional genes catalyzing the anaerobic reduction of nitrite to nitrogen confirmed the occurrence of denitrifiers belonging to the Betaproteobacteria in sponges (Hoffmann et al. 2009).

3. Archaea associated with marine sponges

Once thought to inhabit only extreme environments, Archaea are now known to occur in a wide range of habitats (Stein et al. 1996). In the last years several studies emerged, which describe the detection of Archaea in marine sponges. The first reported archaeon in a sponge was C. symbiosum, a psychrophilic microorganisms affiliated to the crenarchaeal group I.1A, which builds a specific association with the sponge Axinella mexicana (Preston et al. 1996). Whereas C. symbiosum was the only crenarchaeal phylotype inhabiting this sponge, broader association of Crenarchaeota with marine sponges has been demonstrated in other host species (Margot et al. 2002; Lee et al. 2003; Holmes et al. 2007). It has been shown, that members of the group I.1A Crenarchaeota are widespread in marine environments (Karner et al. 2001) and nearly all sponge-derived archaeal sequences are affiliated with this group. The marine isolate Candidatus “Nitrosopumilus maritimus” (Könneke et al. 2005) that grows chemolithoautotrophically by oxidizing ammonia to nitrite, is the only cultivated member of this group. Aerobic ammonia oxidation as energy metabolism for autotrophic growth is also proposed for C. symbiosum. Analysis of the genome sequence of C. symbiosum predicted genes encoding ammonia monooxygenase subunits, ammonia permease, urease, and urea transporters, which implies the use of reduced nitrogen compounds as energy sources for autotrophic growth by ammonia oxidation (Hallam et al. 2006). Furthermore, an autotrophic carbon assimilation cycle with utilization of carbon dioxide as carbon source, as well as an oxidative tricarboxylic acid cycle indicative for additional consumption of organic carbon was supported by genomic analysis.

Although most archaeal sequences retrieved from the host affiliate to the crenarchaeotal lineage, Archaea belonging to the Phylum Euryarchaeota were also reported from marine sponges (Webster et al. 2001; Holmes et al. 2007). Additionally, biomarker analysis as well as fluorescence in situ hybridizations suggested the presence of both Crenarchaeota and Euryarchaeota in the mesohyl of an arctic deep-water sponge (Pape et al. 2006). Whereas for at least some sponge associated Crenarchaeota strong evidence for a functional role in the nitrogen cycle is given, the function of Euryarchaeota remains unclear,

117 although members of this group possess metabolic capabilities of potential benefit to host sponges (e.g. sulfur metabolism).

It was the aim of the third part of this thesis to contribute to a better understanding of these complex sponge-microbe associations by (i) performing a comprehensive meta-analysis of all public available sponge derived 16S rRNA gene sequences supplemented by sequences retrieved from three additional sponges as well as by (ii) addressing the mode of transmission of sponge associated Archaea with a putative role in the nitrogen cycle.

118 4. References

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123

PAPER I

Sponge-Associated Microorganisms: Evolution, Ecology, and Biotechnological Potential

Michael W. Taylor, 1,2 Regina Radax, 1 Doris Steger1 and Michael Wagner1

1Department of Microbial Ecology, University of Vienna, Althanstr.14, A-1090 Vienna, Austria. 2School of Biological Sciences, University of Auckland, Private Bag 92010, Auckland, New Zealand.

Concept by M.W. and M.W.T. R.R. and D.S. made the clone libraries. D.S, M.W.T. and R.R. performed phylogenetic analyses. M.W.T wrote the manuscript.

MICROBIOLOGY AND MOLECULAR BIOLOGY REVIEWS, June 2007, p. 295–347 Vol. 71, No. 2 1092-2172/07/$08.00ϩ0 doi:10.1128/MMBR.00040-06 Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Sponge-Associated Microorganisms: Evolution, Ecology, and Biotechnological Potential† Michael W. Taylor,* Regina Radax, Doris Steger, and Michael Wagner Department of Microbial Ecology, University of Vienna, Althanstr. 14, A-1090 Vienna, Austria

INTRODUCTION ...... 295 EVOLUTION AND DIVERSITY OF SPONGE-ASSOCIATED MICROORGANISMS ...... 297 Known Diversity of Microorganisms from Sponges ...... 298 Existing Evidence for Sponge-Specific Microorganisms ...... 299 Census of Sponge-Associated Microorganisms...... 300 Sponge-Associated Microorganisms: Ancient Partners or Recent Visitors That Have Come To Stay?.....314 Scenario 1: Ancient symbioses maintained by vertical transmission...... 314 Scenario 2: Parental and environmental symbiont transmission ...... 316 Scenario 3: Environmental acquisition...... 316 ECOLOGICAL ASPECTS: FROM SINGLE CELLS TO THE GLOBAL SCALE...... 318 Establishment and Maintenance of Sponge-Microbe Associations ...... 318 Physiology of Sponge-Associated Microorganisms...... 320 Carbon...... 321 Nitrogen...... 321 Sulfur...... 323 Other aspects of microbial metabolism in sponges ...... 323 The Varied Nature of Sponge-Microbe Interactions...... 323 Mutualism/commensalism ...... 323 Microorganisms as a food source for sponges ...... 324 Harming the host: pathogenesis, parasitism, and fouling ...... 325 The Big Picture: Temporal and Biogeographic Variability in Microbial Communities of Sponges...... 326 BIOTECHNOLOGY OF SPONGE-MICROBE ASSOCIATIONS: POTENTIAL AND LIMITATIONS ...... 329 Biologically Active Chemicals from Marine Sponge-Microbe Consortia and Their Commercial-Scale Supply...... 329 Methods for Accessing the Hidden Chemistry of Marine Sponges...... 331 Cultivation of metabolite-producing microorganisms ...... 331 Sponge culture...... 333 Metagenomics...... 334 Cell separation and metabolite localization...... 335 Other Biotechnologically Relevant Aspects of Sponges...... 336 CONCLUSIONS AND FUTURE DIRECTIONS...... 337 ACKNOWLEDGMENTS ...... 337 REFERENCES ...... 338

INTRODUCTION wide variety of marine and freshwater (somewhat more re- stricted) systems and are found throughout tropical, temper- Marine sponges represent a significant component of benthic ate, and polar regions (167). Sponges have been the focus of communities throughout the world, in terms of both biomass much recent interest (Fig. 1) due to the following two main and their potential to influence benthic or pelagic processes (73, 74, 124, 220). Sponges (phylum Porifera) are among the (and often interrelated) factors: (i) they form close associa- oldest of the multicellular animals (Metazoa) and possess rel- tions with a wide variety of microorganisms and (ii) they are a atively little in the way of differentiation and coordination of rich source of biologically active secondary metabolites. This tissues (26, 371). They are sessile, filter-feeding organisms increasing research interest has greatly improved our knowl- which, despite a simple body plan, are remarkably efficient at edge of sponge-microbe interactions, and yet, as apparent obtaining food from the surrounding water (290, 308, 443). throughout this article, many gaps remain in our knowledge of The more than 6,000 described species of sponges inhabit a these enigmatic associations. For example, we still lack a clear picture of microbial diversity—and the factors which influence it—in these hosts. Similarly, the physiology of most sponge- * Corresponding author. Present address: School of Biological Sci- associated microorganisms remains unclear, as do many fun- ences, Faculty of Science, The University of Auckland, Private Bag damental aspects of sponge symbiont ecology. (Throughout 92019, Auckland, New Zealand. Phone: 64 9 373 7599, ext. 82280. Fax: this article, the terms “symbiont” and “symbiosis” are used in 64 9 373 7416. E-mail: [email protected]. † Supplemental material for this article may be found at http://mmbr their loosest possible definitions, to refer simply to two [or .asm.org/. more] different organisms that live together over a long period

127 Part I

FIG. 1. Increasing research interest in marine sponge-microorganism associations. (A) Number of publications retrieved from the ISI Web of Science database by using the following search string: (sponge* or porifera* or demospong* or sclerospong* or hexactinellid*) and (bacteri* or prokaryot* or microbe* or microbial or microorganism* or cyanobacteri* or archaeon or archaea* or crenarchaeo* or fung* or * or dinoflagellate* or zooxanthella*) not (surgery or surgical). (B) Number of sponge-derived 16S rRNA gene sequences deposited in GenBank per year. The 2006 value includes the 184 sequences submitted to GenBank from this article. The search string used to recover sequences was as follows: (sponge* or porifera*) and (16S* or ssu* or rRNA*) not (18S* or lsu* or large subunit or mitochondri* or 23S* or 5S* or 5.8S* or 28S* or crab* or alga* or mussel* or bivalv* or crustacea*). of time, similar to the original de Bary definition. No judgment sponges), Calcarea ( sponges), and Demospongiae is made regarding benefit to either partner.) Here we aim to (), with the last group containing the majority of provide a comprehensive review of the current knowledge of extant species (38, 167). Sponge architecture is unlike that for the evolution, ecology, and biotechnological potential of sponge- any other taxon, and sponge morphology greatly affects many microbe associations. aspects of sponge biology, including interactions with micro- We begin with an introduction to the host organism. The organisms. The basic body plan comprises several different cell phylum Porifera is a paraphyletic grouping consisting of three layers (Fig. 2) (371). The outer surface, or pinacoderm, is major sublineages (classes), namely, the Hexactinellida (glass formed by epithelial cells known as pinacocytes. Through pores

FIG. 2. Schematic representation of a sponge. Arrows indicate the direction of water flow through the sponge. (Adapted from reference 328 with permission of Brooks/Cole, a division of Thomson Learning.)

128 Microorganisms in marine sponges

FIG. 3. Sponges of diverse size, shape, and color. The encrusting sponge Tedania digitata (left), the branching sponge Axinella cannabina (center), and the giant barrel sponge Xestospongia testudinaria (right) are shown. The last two images were kindly provided by Armin Svoboda (Ruhr-Universita¨t, Bochum, Germany).

(ostia) on the sponge surface, these cells also extend along the enemies, such as predators and competitors. Marine sponges interior canals which permeate the sponge. Inside the sponge, have attracted particularly intense scrutiny in this regard, with specialized flagellated cells (choanocytes) form a series of a wide variety of sponge natural products characterized to date chambers where feeding takes place. In these chambers, col- (see reference 32 and its preceding versions). More novel bio- lectively called the choanoderm, the flagellated choanocytes active metabolites are obtained from sponges each year than beat to pump water in through the ostia and along the often from any other marine taxon, and a range of pharmacological elaborate aquiferous systems within the sponge. Choanocytes properties have been demonstrated (32, 250). Various ecolog- also filter out food particles (including bacteria and microal- ical roles have also been proposed for these compounds, in- gae) from the water, and these are transferred to the mesohyl, cluding defense against predators (20, 55, 275), competitors an extensive layer of connective tissue (Fig. 2). In the mesohyl, (94, 395, 411), fouling organisms (363, 487), and microbes (19, food particles are digested via phagocytosis by another group 254, 398). Interestingly, in at least some cases, the compounds of sponge cells, the archaeocytes. These totipotent cells are appear to be produced by associated microorganisms rather capable of differentiating into any of the other sponge cell than by the sponge (27, 285, 351). Continued investigations of types. Also present in the mesohyl of many sponges are dense sponge-derived compounds and their biotechnological and communities of microorganisms (106, 430, 471–473). The ex- ecological implications should guarantee vigorous interest in istence of these putative symbionts alongside bacterium-digest- sponge-microbe associations for some time to come. ing archaeocytes is somewhat paradoxical and implies either Interactions between sponges and microorganisms occur in recognition of different microbial types by the sponge cells or many forms. To a sponge, different microbes can represent shielding of symbiont cells to prevent consumption (482). Once food sources (290, 307, 308), pathogens/parasites (16, 171, 199, filtered in the choanocyte chambers, water is eventually ex- 455), or mutualistic symbionts (474, 477). Microbial associates pelled from the sponge via the exhalant opening, or osculum. can comprise as much as 40% of sponge tissue volume (427), It has been estimated that up to 24,000 liters of water can be with densities in excess of 109 microbial cells per ml of sponge pumped through a 1-kg sponge in a single day (443). tissue (159, 453), several orders of magnitude higher than Beyond the basic body plan described above, sponge mor- those typical for seawater. The diversity in types of interaction phology is highly diverse. Inspection of any marine “sponge is matched by the phylogenetic diversity of microbes that occur garden” will reveal a colorful array of encrusting, branching, within host sponges. It was already evident from early micros- cup-shaped, and massive (amorphous) types (Fig. 3), with in- copy and culturing studies of sponge-associated microbes that dividuals ranging in size from a few millimeters to more than a high levels of morphological and metabolic diversity were meter in diameter (328). Sponge morphology can also reflect present (62, 218, 336, 430, 471–473). The application of mo- ecological function, as seen in the many cyanobacterium-con- lecular tools over the past decade has greatly extended the taining species whose flattened shapes allow optimal light re- known diversity of microorganisms within these hosts (100, ception for their photosynthetic symbionts (337, 474, 477). 106, 146, 214, 294, 390, 458). Each of the three domains of life, Structural integrity is conferred upon most sponges by siliceous i.e., Bacteria, Archaea, and Eukarya, are now known to reside or calcareous spicules (371), and these skeletal components within sponges. We now consider in detail this enormous di- are the basis for much of sponge biology and taxonomy. A wide versity together with the evolutionary mechanisms driving its range of spicule types are secreted, many of which are charac- existence. teristic of particular taxa (167). Collagenous tissues, such as spongin, also play a role in providing structural support and, EVOLUTION AND DIVERSITY OF SPONGE-ASSOCIATED together with spicules, allow the development of very large MICROORGANISMS individuals, such as those found among many tropical species. Sessile organisms such as sponges and other marine inver- Marine sponges are widely considered the most primitive of tebrates (including corals and ascidians) rely heavily on the the metazoans, arising at least as early as the Precambrian, production of chemicals as a form of defense against natural some 600 million years ago (206). According to molecular

129 Part I

clocks, the divergence of sponges from the ancestors of other in a recent 16S rRNA gene library constructed from the fresh- metazoans may have occurred even earlier, around 1.3 billion water sponge Spongilla lacustris (123). Moreover, many of years ago (144). During subsequent periods of the Paleozoic these sequences were highly similar to those known previously era, sponges accounted for much of the biomass on marine from freshwater habitats, suggesting that they may not represent reefs (167, 491). Today, they remain important members of true symbionts. both shallow- and deep-water communities, occupying as much With a few exceptions in the Euryarchaeota (164, 456), ar- as 80% of available surfaces in some areas (74). Such sustained chaea reported from marine sponges are members of the phy- evolutionary and ecological success is probably due, at least in lum Crenarchaeota (164, 200, 226, 294, 454, 456). Lipid biomar- part, to their intimate associations with microbial symbionts. kers also suggested the presence of both crenarchaeotes and However, unlike many other studied host-microbe associa- euryarchaeotes in a deep-water Arctic sponge, though no phy- tions, in which only a very small number of participants are logenetic information was provided in that study (272). The involved (e.g., squid-Vibrio fischeri [258], -Chlamydiae group I.1A Crenarchaeota are extremely prevalent in marine [168], and Bugula-“Endobugula” symbioses [142, 210]), it is environments (180), and almost all sponge-derived archaeal apparent that sponge-associated microbial communities can be sequences are affiliated with this group. The best-studied highly diverse, with a range of different microorganisms con- sponge-associated archaeon is the psychrophilic crenarchaeote sistently associated with the same host species. In this section, “Candidatus Cenarchaeum symbiosum,” which comprises up we describe the extent of this diversity, providing in-depth to 65% of prokaryotic cells within the Californian sponge phylogenetic analyses of all known sponge-associated microor- Axinella mexicana (135, 294, 343, 345). ganisms. We summarize current evidence for the existence of sponge-specific microorganisms and conclude by considering Eukaryotic microbes also occur in sponges. Sponge-inhabit- whether sponge-microbe associations are evolutionarily an- ing dinoflagellates (120, 152, 153, 338, 339, 355, 382, 454, 477) cient or are, instead, recently initiated relationships involving and (16, 47, 51, 53, 65, 113, 305, 390, 409, 454) have microorganisms which are present in the surrounding sea- been reported, with the latter seemingly most prevalent in water. polar regions (16, 51, 53, 113, 305, 409, 454). Freshwater sponges often contain endosymbiotic , primarily zoochlorellae (30, 108, 109, 331, 333, 475, 488). Two previous Known Diversity of Microorganisms from Sponges reports of in sponges were noted by Wilkinson Prior to this review, the diversity of microorganisms (477), while marine sponge-derived fungi are receiving increas- known from sponges was categorized in 14 recognized bacterial ing attention due to their biotechnological potential (44, 163, phyla (and one candidate phylum), both major archaeal lin- 191). Interestingly, of 681 fungal strains isolated worldwide eages, and assorted microbial eukaryotes (145, 148, 477). Se- from 16 sponge species, most belonged to genera which are quences representing the following bacterial phyla have been ubiquitous in terrestrial habitats (e.g., Aspergillus and Penicil- recovered from 16S rRNA gene libraries and/or excised dena- lium) (163). It thus remains unclear in most cases whether such turing gradient gel electrophoresis (DGGE) bands: Acidobac- fungi are consistently associated with the source sponge, or teria, Actinobacteria, Bacteroidetes, Chloroflexi, Cyanobacteria, even whether they are obligate marine species. Compelling Deinococcus-Thermus, Firmicutes, Gemmatimonadetes, Nitro- evidence for symbiosis of a yeast with sponges of the genus spira, Planctomycetes, Proteobacteria (Alpha, Beta, Delta, and Chondrilla was obtained by extensive microscopy studies of Gammaproteobacteria), Spirochaetes, and Verrucomicrobia (7, both adult sponge tissue and reproductive structures, with 95, 123, 146, 148, 151, 154, 214, 317, 342, 383, 390, 391, 396, strong indications of vertical transmission of the yeast symbi- 404, 407, 421, 452, 454, 458; S. R. Longford, N. A. Tujula, G. R. ont (221). Crocetti, A. J. Holmes, C. Holmstro¨m, S. Kjelleberg, P. D. Little is known about viruses in sponges, although virus-like Steinberg, and M. W. Taylor, unpublished data). In addition, a particles were observed in cell nuclei in Aplysina (Verongia) seemingly sponge-specific candidate phylum, “Poribacteria,” cavernicola (432). It was suggested that these particles could be has also been reported for several sponges (100). The most involved in sponge cell pathology. Infection of an Ircinia stro- frequently recovered sequences in general 16S rRNA gene bilina-derived alphaproteobacterium by a bacteriophage iso- surveys of sponges include those from the Acidobacteria, Acti- lated from seawater has also been demonstrated (211), al- nobacteria, and Chloroflexi (148). Members of several bacterial though the propensity of this siphovirus to infect the bacterium phyla, namely, the Actinobacteria, Bacteroidetes, Cyanobacteria, Firmicutes, Planctomycetes, Proteobacteria, and Verrucomicro- in nature is not known. bia, have also been isolated in pure culture from marine In addition to the realization of high microbial diversity per sponges (46, 47, 56, 81, 95, 147, 187, 188, 198, 202, 214, 235, se, we are now beginning to recognize more subtle patterns of 263, 264, 292, 334, 341, 365, 453, 458). Sequences from the host-symbiont distribution. For example, it appears that a Chlorobi (green sulfur bacteria) have not been obtained from given species of sponge contains a mixture of generalist and sponges, although positive fluorescence in situ hybridization specialist microorganisms (390) and that the associated micro- (FISH) signals were obtained from Rhopaloeides odorabile with bial communities are fairly stable in both space and time (105, a specific probe for this phylum (458). In contrast to the case 390, 391, 454). One particularly interesting pattern to emerge for marine sponges, the (limited) available evidence for fresh- is the apparent widespread existence of sponge-specific bacte- water species suggests that bacterial diversity and abundance rial clusters, i.e., closely related groups of bacteria which are are both much lower. Only sequences from the Actinobacteria, found only in sponges (146). In the following section, we ex- Chloroflexi, and Alpha- and Betaproteobacteria were recovered amine the published evidence for such clusters.

130 Microorganisms in marine sponges

Existing Evidence for Sponge-Specific Microorganisms most cases, were strongly supported by bootstrap analyses (in all cases, the clusters were found with three different tree The notion that marine sponges might contain a specific construction methods). Three further clusters—each sponge microbiota arose some 3 decades ago from the seminal work of specific, with the exception of a single nonsponge sequence— Vacelet et al. and Wilkinson et al. (427, 430, 469, 471–473, were also identified in the Acidobacteria and in a lineage of 483). Based on electron microscopy and bacterial cultivation uncertain affiliation (later recognized as Gemmatimonadetes studies, these pioneers of sponge symbiont research proposed (146, 499). Overall, 70% of the 190 sponge-derived sequences the following three broad types of microbial associates in available at the time fell into one of these monophyletic clus- sponges: (i) abundant populations of sponge-specific microbes ters or the other. Interestingly, within-cluster 16S rRNA se- in the sponge mesohyl, (ii) small populations of specific bac- quence similarities ranged down to as low as 77% (146), often teria occurring intracellularly, and (iii) populations of nonspe- considered indicative of phylum-level differences (170). Sev- cific bacteria resembling those in the surrounding seawater eral subsequent, mostly cultivation-independent studies have (427, 472). One type of bacterial isolate, regarded as a single also led to the recovery of apparently sponge-specific se- species, was recovered from 35 taxonomically diverse sponges quences. Approximately 50% of 16S rRNA gene sequences in from several geographic regions, but never from seawater (469, a gene library obtained from the unidentified Indonesian 483). Immunological experiments in which these same isolates sponge 01IND 35 were most closely related to sequences de- cross-reacted with other “sponge-specific” bacteria but not rived from other sponges (154). These included members of with seawater isolates were taken as further evidence of sponge the Acidobacteria, Nitrospira, Bacteroidetes, and Proteobacteria, specificity (469). Another significant advance came in 2002, as well as several sequences in a group of uncertain affiliation when Hentschel and coworkers integrated these concepts into (our analyses indicate that these may be deltaproteobacterial the molecular age (146). They defined sponge-specific clusters sequences [see Fig. 8]). A similar situation was reported for as sponge-derived groups of at least three 16S rRNA gene Discodermia dissoluta, whereby three-quarters of 160 retrieved sequences which (i) are more similar to each other than to 16S rRNA sequences were most similar to other sponge-de- sequences from other, nonsponge sources; (ii) are found in at rived sequences (342). Conversely, of 21 unique sequences least two host sponge species and/or in the same host species (each representing a unique restriction fragment length poly- but from different geographic locations; and (iii) cluster to- morphism [RFLP] type) obtained from the Caribbean sponge gether irrespective of the phylogeny inference method used Chondrilla nucula, only 5 retrieved other sponge-derived 16S (146). rRNA sequences during BLAST searches (although with the The hypothesis of widespread, sponge-specific microbial advantage of our larger data set, we found indications that communities put forward by Hentschel and colleagues (146) several more of the C. nucula sequences are in fact from was compelling and was constrained only by the limited data members of sponge-specific clusters) (151). Perhaps the most set available at that time. They performed phylogenetic anal- impressive sponge-specific cluster to be reported so far is the yses with the 190 publicly available sponge-derived 16S rRNA candidate phylum “Poribacteria” (100). Fieseler and colleagues gene sequences, the majority of which were from Aplysina found members of this lineage, which is moderately related to aerophoba, Rhopaloeides odorabile, and Theonella swinhoei. the Planctomycetes, Verrucomicrobia, and Chlamydiae (446), in These three sponges are phylogenetically only distantly related several sponges from geographically diverse locations, but and were collected from the Mediterranean Sea, the Great never in adjacent seawater or sediment samples (100). It will Barrier Reef, and Micronesia/Japan/Red Sea, respectively, yet be especially interesting to see whether “Poribacteria” se- they contained largely overlapping microbial communities. To- quences are recovered from other environments in the future. gether with the earlier work of Wilkinson and contemporaries The sheer number of reports dealing with sponge-specific (e.g., see reference 483), these remarkable results suggested microorganisms is compelling. However, one must be cautious that even unrelated sponges with nonoverlapping geographic when proposing a sponge-specific cluster. Of crucial impor- ranges might share a common core of bacterial associates. tance is the selection of nonsponge reference organisms for Indeed, subsequent studies have lent further weight to this phylogenetic analyses. In principle, any group of sequences can notion, with numerous reports of similar (and in some cases appear sponge specific if the most appropriate reference or- sponge-specific) bacteria found in different sponge species ganisms (i.e., those that are most closely related to the sponge- (100, 151, 154, 198, 235, 342, 404, 407). Furthermore, both derived sequences) are not also included. The length of analyzed cultivation-based and molecular methods have provided evi- sequences is also of concern, with the level of phylogenetic dence for distinct microbial communities between sponges and information obtainable increasing with sequence length. Every the surrounding seawater (151, 265, 334, 391, 472). Taken effort should be made to obtain at least one near-full-length together, these results appear to indicate that sponge-associ- sequence per sequence type (or operational taxonomic unit). ated microbial communities are indeed unique and at least Decreasing sequence costs render this eminently achievable, partially sponge specific, and the existence of sponge-specific and in many cases, it would only involve performing a few microorganisms has consequently become something of a par- additional sequencing reactions. These are not new ideas adigm in this field. and we are certainly not the first to advocate the use of A total of 14 monophyletic, sponge-specific sequence clus- full-length sequences (e.g., see reference 216), but during ters were identified in the original study of Hentschel et al. our analyses of sponge-derived 16S rRNA sequences, it be- (146). These occurred in the Acidobacteria, Actinobacteria, came apparent that many of these sequences are rather Bacteroidetes, Chloroflexi, Cyanobacteria, Nitrospira, and Pro- short and therefore phylogenetically not particularly infor- teobacteria (Alpha, Delta, and Gammaproteobacteria) and, in mative. Indeed, we encountered many problems with inser-

131 Part I

FIG. 4. 16S rRNA-based phylogeny showing representatives of all bacterial and archaeal phyla from which sponge-derived sequences have been obtained. Sponge-derived sequences are shown in bold, with additional reference sequences also included. The displayed tree is based on a maximum likelihood analysis. Bar, 10% sequence divergence.

tion of short sponge-derived sequences into our phyloge- to provide an overview of microbial diversity in sponges and netic trees, and in some cases, we were not even certain of (ii) to critically assess the occurrence of monophyletic, sponge- their phylum-level affiliation. specific sequence clusters. As mentioned above, such clusters are often discussed, yet their existence has not been reevalu- ated rigorously in light of the rapidly expanding 16S rRNA Census of Sponge-Associated Microorganisms sequence databases. It is thus unclear whether these clusters Increasing interest in sponge-microbe associations has re- are truly sponge specific or merely reflect a greater sampling sulted in a concomitant increase in the amounts of 16S rRNA effort for these communities than for others. sequence data obtained from sponges (Fig. 1B). There are We began, using the ARB program package (217), by estab- currently ϳ1,500 sponge-derived 16S rRNA gene sequences lishing an encompassing database that contains all sponge- available in GenBank (http://www.ncbi.nlm.nih.gov/), in con- derived 16S rRNA sequences which were available in GenBank trast to only 190 such sequences available for the 2002 study by on 28 February 2006. In addition to these 1,499 sequences Hentschel et al. (146). We carried out an extensive phyloge- (plus 11 18S rRNA sequences amplified from eukaryotic mi- netic analysis of all currently available sponge-derived 16S crobes in sponges), we contributed a further 184 bacterial and rRNA gene sequences, with two main objectives, as follows: (i) archaeal sequences from three hitherto unstudied sponges,

132 Microorganisms in marine sponges

TABLE 1. Summary of all publicly available sponge-derived 16S rRNA sequence data (as of 28 February 2006) plus 184 bacterial and archaeal sequences contributed from this article

No. of sequences % of sequences in clusters obtained from:

Total no. of No. of sequences Sponges and one Phylogenetic affiliation Obtained from Exclusively Sponges and sequences of Ͼ1,000 bp Uncultivated nonsponge an isolate spongesa corals organism Bacteria 1,630 592 Acidobacteria 66 9 66 0 5 64 24 Actinobacteria 266 99 190 92 38 (45) 0 1 Bacteroidetes 77 20 46 31 0 (4) 0 10 Chloroflexi et al. 109 48 109 0 62 (75) 0 0 Cyanobacteria 119 68 111 7 79 0 0 Deinococcus-Thermus 20 2000 0 Firmicutes 96 31 45 51 9 0 0 Gemmatimonadetes 16 5 16 0 25 56 0 Lentisphaerae 11 1000 0 Nitrospira 14 6 14 0 57 29 0 Planctomycetes 11 4 9 2 0 0 0 “Poribacteria” 21 10 21 0 100 0 0 Proteobacteria Alpha- 311 125 196 115 14 (22) 0 4 Beta/Gamma- 430 114 298 134 34 (37) 0 0 Delta- 48 25 48 0 15 40 6 Spirochaetes 62 60670 0 TM6 1 1 1 0 0 0 0 Verrucomicrobia 13 13 12 1 23 0 0 Uncertain affiliation 23 11 23 0 78 0 0

Archaea 44 10 Crenarchaeota 43 10 43 0 28 (42) 0 0 Euryarchaeota 10 1000 0

Eukaryab 20 6 18 2

Total 1,694 608 1,259 435 546 74 43

a Numbers in parentheses are inclusive of clusters which are supported by only two of three tree construction methods. b Includes both 18S rRNA- and 16S rRNA (plastid)-derived sequences.

namely, Agelas dilatata, Plakortis sp. (both from the Bahamas; mus, Firmicutes, Gemmatimonadetes, Nitrospira, Planctomycetes, kindly provided by U. Hentschel), and Antho chartacea (from “Poribacteria,” Proteobacteria [Alpha-, Beta-, Delta-, and Gam- southeastern ). Preliminary phylogenetic analyses maproteobacteria], Spirochaetes, Verrucomicrobia, and Chlorobi identified members of putative sponge-specific clusters, and for FISH signals), we report for the first time the presence in each cluster, the most similar nonsponge sequences were re- sponges of 16S rRNA sequences affiliated with the phylum trieved by BLAST searches (from both regular NCBI and Lentisphaerae and the candidate phylum TM6. The number of environmental genome databases) and imported into ARB for sequences representing each phylum varied widely, from single subsequent alignment (automatic and manual). The resulting sequences for the Lentisphaerae and TM6 to more than 250 ARB database, containing an alignment of all sponge-derived sequences for each of the Actinobacteria, Alphaproteobacteria, sequences and their nearest relatives (together with annotated and Beta/Gammaproteobacteria (Table 1). The proportions of information [e.g., host species and collection location] for the sequences derived from cultivated versus noncultivated micro- sponge sequences), is available upon request. Extensive phy- organisms also varied greatly among phyla. logenetic analyses (see the supplemental material for details) The phylogenetic analyses presented here strongly support were conducted, taking all (n ϭ 1,694) sponge-derived se- the existence of monophyletic, sponge-specific 16S rRNA se- quences into account. In order to rigorously test the existence quence clusters. These occurred in many of the bacterial and of monophyletic, sponge-specific sequence clusters, we em- archaeal phyla found in sponges, with approximately one-third ployed multiple tree construction methods (maximum likeli- (32%) of all sponge-derived sequences falling into such clus- hood, neighbor joining, and maximum parsimony), together ters (Table 1; Fig. 5 to 15; also see the supplemental material). with the use of various sequence conservation filters and cor- If sequences derived from cultured isolates are excluded, this rection parameters. figure rises to 42%. This result was expected since tightly In total, sequences representing 16 bacterial phyla and both linked symbionts—those presumed to occur in sponge-specific major archaeal lineages (Crenarchaeota and Euryarchaeota) clusters—are likely difficult to cultivate and therefore under- have been recovered from sponges (Fig. 4). In addition to represented in culture collections. Several additional clusters those known prior to this study (Acidobacteria, Actinobacteria, each contained a single nonsponge sequence, with the extra Bacteroidetes, Chloroflexi, Cyanobacteria, Deinococcus-Ther- sequences often, but not always, obtained from marine envi-

133 Part I

FIG. 5. 16S rRNA-based phylogeny of sponge-associated cyanobacteria and . Sponge-derived sequences are shown in bold. The displayed tree is a maximum likelihood tree constructed based on long (Ն1,000 nucleotides) sequences only. Shorter sequences were added using the parsimony interactive tool in ARB and are indicated by dashed lines. Shaded boxes represent sponge-specific monophyletic clusters, as defined

134 Microorganisms in marine sponges

ronments. It is also possible that sponge-specific microbes are study of Hentschel et al. (146), no doubt reflecting the fact that more prevalent in those sponges which contain very dense most of the relevant coral sequences were deposited in GenBank microbial communities (Ute Hentschel, personal communica- since then. It is too early to speculate whether some sort of tion), i.e., the so-called bacteriosponges or high-microbial- marine invertebrate-specific sequence cluster exists, but fur- abundance sponges (148, 430). Due to a lack of microbial ther sampling of taxa such as ascidians and bryozoans should abundance data for most host sponges, we did not attempt to help to resolve this issue. A study of two marine macroalgae take this factor into account during our analyses. Overall, while and the cooccurring sponge Cymbastela concentrica gave no representation of sequences in sponge-specific clusters was indication of specific clusters spanning these taxonomically quite high, it should be noted that the proportions of se- disparate groups (Longford et al., unpublished data). quences falling within such clusters differed greatly among the Sponge-specific sequence clusters were not prevalent for, various phyla. among others, the Bacteroidetes (see Fig. S1 in the supplemen- More than three-quarters of the 119 available sponge-de- tal material) and Firmicutes (see Fig. S2 in the supplemental rived Cyanobacteria sequences fell into monophyletic, sponge- material), perhaps reflecting the relatively high proportions of specific clusters (Table 1; Fig. 5). Most of these were in two sequences derived from cultivated organisms in these phyla. clusters, with one comprising 25 sequences from at least 7 We report for the first time the recovery of Lentisphaerae sponge species and the other comprising 52 sequences from 21 (Fig. 11) and candidate phylum TM6 (Fig. 12A) sequences sponges. The latter cluster represents the recently described from sponges. Each phylum was represented by a single 16S candidate species “Candidatus Synechococcus spongiarum” rRNA sequence, from the marine sponges Plakortis sp. and (426) and was the sole Cyanobacteria cluster in the study of Antho chartacea, respectively, and it cannot be ruled out that Hentschel et al. (146), while the former corresponds to the these represent contaminating sequences from the sur- filamentous cyanobacterium Oscillatoria spongeliae (39, 157). rounding environment (although arguably this also applies Sequences representing O. spongeliae were not available for to many, more commonly recovered sequence types). The the 2002 study. Several other, smaller clusters are also evident Lentisphaerae phylum comprises part of the so-called Planc- among the cyanobacteria (Fig. 5). Additionally, in a number of tomycetes-Verrucomicrobia-Chlamydiae (PVC) superphylum cases, microalgal plastids have also been amplified using 16S (446), with sponge-derived sequences from the superphylum rRNA primers. additionally being found in the Verrucomicrobia, Planctomyce- Another bacterial phylum containing many sponge-specific tes, and “Poribacteria” (Fig. 11). Members of the superphylum sequence clusters is the Chloroflexi (Table 1; Fig. 6). Of the 109 are frequently associated with eukaryotes. There is also a sponge-derived sequences analyzed, 62% comprised such clus- group of uncertain affiliation which falls near the PVC super- ters, while the occurrence of a further 13% of sequences in phylum (but without strong bootstrap support) during phylo- clusters was weakly supported. In the new analyses, all but one genetic analyses. This group includes sequences from many of the members of a sponge-specific cluster described by sponges, such as Agelas dilatata, Aplysina aerophoba, Discoder- Hentschel and coworkers (146) remained in a cluster, although mia dissoluta, and Theonella swinhoei. Those sequences most these sequences were now dispersed over four different clus- closely related to the sponge sequences are also from marine ters. Such movement of sequences was frequently observed environments. and is not surprising given the much larger data set at our Several large sponge-specific clusters were found among the disposal now (i.e., many new related sequences, both sponge- Actinobacteria sequences, particularly in the family Acidimicro- and non-sponge-derived, were included in the phylogenetic biaceae (Fig. 13). The largest comprised 54 sequences obtained analyses described here). None of the sponge-derived se- from sponges from the Caribbean (Agelas dilatata, Discodermia quences were closely related to the few described Chloroflexi dissoluta, Plakortis sp., and Xestospongia muta), Indonesia species, although many were similar to sequences from uncul- (Xestospongia testudinaria), the Red Sea (Theonella swinhoei), tivated organisms, particularly from marine environments and the South China Sea (Dysidea avara). None of the se- (Fig. 6). quences within this cluster were obtained from cultured bac- Interestingly, many sponge-derived 16S rRNA sequences teria, with the nearest (but still distantly related) cultured ac- formed exclusive monophyletic clusters with sequences ob- tinobacteria being the wastewater bacterium Microthrix tained from corals (Table 1). This was particularly apparent for parvicella and the acidophilic Acidimicrobium spp. (Fig. 13). the Acidobacteria and Deltaproteobacteria (Fig. 7 and 8, respec- Although not representing a sponge-specific cluster, the tively) but was also evident for the Gemmatimonadetes (Fig. 9) group of sequences affiliated with the marine Pseudovibrio spp. and Nitrospira (Fig. 10). No coral-derived sequences shared within the Alphaproteobacteria deserves special mention (Fig. monophyletic clusters with sponge sequences in the original 14). Members of this genus are frequently found in sponge-

by Hentschel et al. (146), i.e., a group of at least three sponge-derived 16S rRNA gene sequences which (i) are more similar to each other than to sequences from other, nonsponge sources, (ii) are found in at least two host sponge species and/or in the same host species but from different geographic locations, and (iii) cluster together irrespective of the phylogeny inference method used (all clusters shown here also occurred in neighbor-joining and maximum parsimony analyses). Names outside wedges of grouped sequences represent the sponges from which the relevant sequences were derived; the number in parentheses indicates the number of sequences in that wedge. Filled circles indicate bootstrap support (maximum parsimony, with 100 resamplings) of Ն90%, and open circles represent Ն75% support. The outgroup (not shown) consisted of a range of sequences representing several other bacterial phyla. Bar, 10% sequence divergence.

135 Part I

FIG. 6. 16S rRNA-based phylogeny of sponge-associated Chloroflexi organisms. Details are the same as those provided for Fig. 5, with the following additions. Shaded boxes contained within dotted lines represent sponge-specific clusters supported by only two tree construction methods (ML, maximum likelihood; MP, maximum parsimony; and NJ, neighbor joining), and new sequences from our laboratory have the prefix “AD” (for the sponge Agelas dilatata), “AnCha” (Antho chartacea), or “PK” (Plakortis sp.). derived cultivation-based and molecular studies (95, 96, 147, the fact that almost all of these were within the group I.1A 187, 198, 263, 453), and there is strong evidence for its being a Crenarchaeota bears testimony to their high degree of phylo- true sponge symbiont (95, 453). genetic relatedness. The recently isolated ammonia-oxidizing Only 28% of sponge-derived Archaea sequences fell into archaeon “Candidatus Nitrosopumilus maritimus” (192) is the well-supported sponge-specific clusters (Fig. 15), although only cultivated member of this group, with the well-studied

136 Microorganisms in marine sponges

FIG. 7. 16S rRNA-based phylogeny of sponge-associated Acidobacteria organisms. Details are the same as those provided for Fig. 5 and 6, with the following two additions. Open boxes represent monophyletic clusters containing sponge-derived sequences and a single, nonsponge origin sequence, and open boxes with asterisks outside them signify clusters containing only sponge- and coral-derived sequences (the number of asterisks corresponds to the number of coral-derived sequences within the cluster).

137 Part I

FIG. 8. 16S rRNA-based phylogeny of sponge-associated Deltaproteobacteria organisms. Details are the same as those provided for Fig. 5 to 7.

(but still uncultivated) archaeon “Ca. Cenarchaeum symbio- been recovered from sponges, all of which were marine sum” being the best known sponge-associated member. A ge- sponges (Table 1; Fig. 15). All but one of these was from the nome project for the latter has recently been completed (134). Crenarchaeota, with a single Euryarchaeota sequence from the At the time of sequence collection, 44 archaeal sequences had Great Barrier Reef sponge Rhopaloeides odorabile (456). An

138 Microorganisms in marine sponges

FIG. 9. 16S rRNA-based phylogeny of sponge-associated Gemmatimonadetes organisms. Details are the same as those provided for Fig. 5 to 7. article which appeared in mid-2006 (whose sequences were not are consistent with the earlier study by Hentschel et al. (146), available on 28 February 2006 and were therefore not included with, for example, the Actinobacteria, Nitrospira, and Acidobac- in our study) reported more euryarchaeotal sequences from teria still well represented by sponge-specific microorganisms. various sponges, although the majority of sequences in that As could be expected, some sponge-specific clusters from the study were still affiliated with the Crenarchaeota (164). 2002 study now form parts of several new clusters, while others All sponge-derived 16S rRNA sequences available on 28 do not comprise clusters at all in the new data set. Conversely, February 2006 were analyzed phylogenetically, but for practi- the addition of more sequences meant that many formerly cal reasons the larger trees are available only in the supple- single sequences are now in specific clusters with other sponge- mental material. Broadly speaking, the results of our analyses derived sequences.

FIG. 10. 16S rRNA-based phylogeny of sponge-associated Nitrospira organisms. Details are the same as those provided for Fig. 5 to 7.

139 Part I

FIG. 11. 16S rRNA-based phylogeny of sponge-associated Verrucomicrobia, Planctomycetes, Lentisphaerae, and “Poribacteria” organisms and of a lineage of uncertain affiliation. These and associated lineages comprising the PVC superphylum (446) are shown. Details are the same as those provided for Fig. 5 to 7.

140 Microorganisms in marine sponges

FIG. 12. 16S rRNA-based phylogeny of sponge-associated members of the candidate phylum TM6 (A), Deinococcus-Thermus organisms (B), and Spirochaetes organisms (C). Details are the same as those provided for Fig. 5 to 7. (B) In our analyses, the position of clone Dd-spU-11 (from the sponge Discodermia dissoluta) was not stable, and we are not certain of its phylogenetic affiliation.

Very few sequences were available from sponge-associated we failed to include some key sequences which would have eukaryotic microbes at the time of database establishment broken up otherwise specific sponge clusters. Another factor (since then, some 45 18S rRNA sequences derived from relates to the short lengths of many sponge-derived 16S rRNA sponge-associated fungi have been deposited in GenBank). sequences. We constructed our trees using only sequences Those that are included in our database include 9 16S rRNA longer than 1,000 bp, but more than two-thirds of all sponge- sequences derived from diatom chloroplasts (Fig. 5) and 11 derived sequences are shorter than this (Table 1), and we 18S rRNA sequences obtained from diatoms and dinoflagel- added these via the parsimony interactive tool in ARB. In lates. All but one of the 18S rRNA sequences were obtained principle, this method allows the insertion of short sequences from Antarctic sponges (454), with the remaining sequence without changing tree topology (217). However, when many representing a zooxanthella (Symbiodinium sp.) from the Pa- short sequences are added at once, they can influence each lauan sponge Haliclona koremella (49). other’s positioning (and potentially bias the analysis towards We endeavored to be as thorough and as careful as possible the formation of sponge-specific clusters). We attempted to throughout our analyses, yet there remain some caveats to our gauge the severity of this problem by (for a selection of se- results. Despite extensive BLAST searches using members of quences) sequentially adding and removing individual short all putative sponge-specific clusters, it is not inconceivable that sequences and comparing their placement to the outcome

141 Part I

FIG. 13. 16S rRNA-based phylogeny of sponge-associated Actinobacteria organisms belonging to the family Acidimicrobiaceae. Other sponge- derived actinobacteria are shown in Fig. S3 in the supplemental material. Details are the same as those provided for Fig. 5 to 7.

when they were all added at once. The results were highly tance of obtaining at least one near-full-length sequence for consistent, but it should not be assumed that this will always be each operational taxonomic unit obtained. This is not possible the case. The alternative is to perform the entire phylogenetic in some cases (e.g., excised DGGE bands) but is feasible in analysis with short sequences and to truncate longer sequences many others. to leave only the homologous region; this results in the loss of It is prudent to consider whether the apparent occurrence of much phylogenetic information and is not recommended un- sponge-specific sequence clusters could have a more dubious der any circumstances (216). Again, we reiterate the impor- origin, namely, laboratory contamination. Theoretically, a 16S

142 Microorganisms in marine sponges

FIG. 14. 16S rRNA-based phylogeny of sponge-associated Alphaproteobacteria organisms affiliated with the genus Pseudovibrio and its relatives. Other sponge-derived alphaproteobacteria are shown in Fig. S4 and S5 in the supplemental material. Details are the same as those provided for Fig. 5 to 7.

143 Part I

FIG. 15. 16S rRNA-based phylogeny of sponge-associated archaeal organisms. Details are the same as those provided for Fig. 5 to 7.

144 Microorganisms in marine sponges

rRNA gene-containing plasmid or PCR product could, if ac- Ecology, Vienna, Austria, 20 to 25 August 2006). The lack of cidentally spread to from several sponges in the same new phyla in these data allow one to speculate that the major- laboratory, appear to form its own sponge-specific cluster. ity of sponge-associated microorganisms may have already However, the available evidence strongly suggests that this is been encountered in gene libraries, at least at the phylum level. not the case, since many or most clusters contain sequences However, two major caveats exist. Although we may arguably originating from several independent laboratories. be nearing the point of diminishing returns with respect to With almost 1,700 sponge-derived 16S rRNA sequences fall- using current techniques to recover novel lineages from ing into some 16 or more bacterial and archaeal phyla, we sponges (i.e., gene libraries constructed using general 16S sought to address the following question: how well sampled are rRNA primers), it is highly likely that the use of phylum- marine sponge-associated microbial communities? If current specific primers and/or metagenomic (i.e., PCR-independent) studies are recovering mainly sequences which were previously approaches will reveal phyla previously unknown to exist in obtained from sponges (as the presence of sponge-specific these hosts or even unknown to science (e.g., “Poribacteria”) clusters might imply), then we may have already uncovered (100). To our knowledge, there is no example of a sponge for most of the microbial diversity in these hosts, suggesting which the results of general versus specific 16S rRNA gene that our current descriptive phase might be nearing its log- libraries have been compared. A second point is that few gene ical conclusion. Unfortunately, the available data are insuf- libraries, including those from sponges, are sequenced to full ficient to satisfactorily address this issue for sponges. In a coverage, and it is possible that the recurring sequences ob- recent article in this journal, Schloss and Handelsman (348) tained from different sponges are merely those that are most used the program DOTUR to estimate richness at different abundant (or those that PCR is most biased toward) in each levels of phylogenetic relatedness for each bacterial phylum sponge, with the unsequenced remainder of the library poten- represented in the Ribosomal Database Project (61). To per- tially contributing new sequence types. The advent of high- form an analogous study with the sponge symbiont data set, we throughput sequencing technologies (e.g., see reference 227) were restricted to sequences which met the following criteria: offers the potential to sequence gene libraries to much greater (i) they were part of attempts at extensive microbial commu- depth, illuminating the rare biosphere within sponges (376). nity surveys using general 16S rRNA gene primers for the Statistical comparisons of microbial community composi- construction of clone libraries; (ii) they overlapped a sufficient tions allow for the inclusion of more sequences (relative to distance to be useful ( positions 100 to 500 species richness estimates via DOTUR) due to less stringent would have been appropriate for a reasonable portion of the selection criteria. We thus used the so-called parsimony test, sponge data set); and (iii) they were not obtained from pre- implemented in the program TreeClimber (347), to compare screened gene libraries (e.g., by RFLP analysis), as this would our three new gene libraries (from the sponges Agelas dilatata, heavily bias results—thus, all collected sequences must have Antho chartacea, and Plakortis sp.) with selected sponge- been deposited in GenBank. After applying these (in our eyes) derived libraries from the literature and those deposited in minimal criteria, only 317 sequences (of 1,694) were deemed GenBank. The parsimony test compares phylogenetic trees suitable for use with DOTUR or similar programs. For many rather than sequence data per se (228, 347), and various tree phyla, only a few sequences were retained (e.g., for Cyanobac- construction algorithms can be employed. Our criteria for se- teria, 8 of 119 sequences were kept, and for Alphaproteobacte- quence inclusion were that (i) general 16S rRNA gene primers ria, 21 of 311 sequences were kept), precluding meaningful were used and (ii) at least 25 sequences were available from analyses. Furthermore, even if 50 or more sequences were each library. The main caveats are that prescreening of clones suitable (as in the case of the Beta/Gammaproteobacteria), (e.g., by RFLP analysis) with subsequent representation of these were not necessarily representative of the known each operational taxonomic unit by a single sequence prevents (sponge-derived) diversity within that phylum, again prevent- strict application of the parsimony test (347), while low se- ing the drawing of meaningful conclusions. Although statisti- quencing coverage of some libraries may obscure true similar- cally robust analyses are therefore not possible at this stage, ities or differences among libraries by missing overlapping or data from two recent studies can add greatly to this discussion. distinct sequences, respectively. With these considerations in In the first, Lopez and colleagues at the Harbor Branch Ocean- mind, we compared the three libraries obtained from this study ographic Institution (213) obtained more than 700 sequences with those from the marine sponges Theonella swinhoei (146), from 20 different sponge-derived gene libraries by using gen- Aplysina aerophoba (146), Rhopaloeides odorabile (458), Cym- eral 16S rRNA primers, with the vast majority of these belong- bastela concentrica (Longford et al., unpublished data), Disco- ing to phyla already obtained from sponges, such as Chloroflexi, dermia dissoluta (342), and Chondrilla nucula (151) and the Cyanobacteria, Nitrospira, Planctomycetes, and Spirochaetes.Of freshwater sponge Spongilla lacustris (123). An initial analysis the recovered sequences, was the only comprising all 10 libraries yielded a highly significant (P Ͻ major taxonomic group not previously obtained from sponges. 0.001) result (i.e., the differences in sequence composition In another study, examining the Adriatic sponges Chondrilla among the various libraries were not due to chance). Likewise, nucula and Tethya aurantium, Thiel and coworkers recov- comparisons of the marine versus freshwater (S. lacustris) li- ered representatives of only known sponge-associated phyla braries, as well as comparisons among the marine libraries and (Acidobacteria, Actinobacteria, Bacteroidetes, Cyanobacteria, Gem- among broad geographic locations, were all highly significant. matimonadetes, Planctomycetes, Proteobacteria [Alpha-, Beta-, The usefulness of such analyses should increase as more 16S Delta-, and Gammaproteobacteria], Spirochaetes, and Verru- rRNA gene libraries are sequenced from sponges (and with comicrobia) (404; V. Thiel, T. Staufenberger, and J. F. Imhoff, greater sequencing coverage), including multiple species from presented at the 11th International Symposium on Microbial the same location and/or from the same genus or family.

145 Part I

FIG. 16. Summary of various evolutionary scenarios for sponge-microorganism associations.

Sponge-Associated Microorganisms: Ancient Partners plexity of microbial communities in seawater could have led to or Recent Visitors That Have Come To Stay? this bacterium being missed in Wilkinson’s culture libraries. In the 22 years since the Wilkinson study, a wealth of mo- Based largely upon arguments centering on immunological lecular data has become available for sponge-associated mi- evidence dating back to the 1980s (469), it is often stated that croorganisms, ranging from sequences of single genes to an sponge-bacterium symbioses have existed for as long as 600 entire genome (for the archaeon “Candidatus Cenarchaeum million years. This would date such associations back to the symbiosum”) (134). Here we ponder whether such data can be Precambrian, prior to the bulk of taxonomic radiation in exploited to address the issue of the ancientness (or otherwise) sponges. Moreover, given the likely basal position of sponges of sponge-microbe associations. First, we consider some of the in the metazoan phylogenetic tree (38, 133), this would pre- many possible evolutionary scenarios (summarized in Fig. 16), sumably make sponges and microorganisms the most ancient as follows. of all metazoan-microorganism associations. So what is the Scenario 1: Ancient symbioses maintained by vertical trans- evidence for this oft-cited ancient symbiosis? In his 1984 study, mission. A given sponge-specific cluster in the phylogenetic on which the majority of these arguments rest, Wilkinson used tree of life may contain 16S rRNA gene sequences derived a collection of 296 sponge isolates which, on the basis of mor- from distantly related, geographically disparate sponge species. phological and physiological characteristics, comprised one If the microorganisms represented by these sequences do not bacterial species (469). In addition, 128 seawater and nonspe- occur outside sponges today, then the ancestral strain (the cific sponge isolates were included as control strains. It is future symbiont) may have first inhabited a sponge during one important to note that the sponge-specific isolates were ob- or several colonization events prior to sponge speciation (ϳ600 tained from phylogenetically distant sponges in widely sepa- million years ago) (the Precambrian acquisition hypothesis of rated geographical regions. From seven of the specific isolates Wilkinson [469]). Such a symbiosis could have been main- and five of the others, Wilkinson prepared antisera and per- tained in the intervening years via vertical transmission (see formed agglutination tests. Many of the “sponge-specific” “Establishment and Maintenance of Sponge-Microbe Associ- strains reacted positively in these tests to one or more of the ations”), and the microbes evolved to become sponge (or even antisera derived from sponge-specific bacteria, but none of species) specific. A related but subtly different hypothesis is them reacted with sera derived from non-sponge-specific that an association could still be ancient but not predate the strains, nor did cross-reactions occur between the 128 non- bulk of sponge speciation. In this scenario, it is conceivable sponge-specific strains and the sera prepared from sponge- that one sponge could have been colonized very early on, specific bacteria. The implication of these results was that the resulting in the evolution of a sponge-adapted microorganism. studied, widespread, sponge-specific bacterium did indeed Millions (or hundreds of millions) of years later, this microbe form a single species group distinct from isolates found in the could have spread across the oceans and, upon encountering surrounding seawater (469). According to Wilkinson, the most other sponges, colonized them. Perhaps it is no longer present logical explanation for the occurrence of this specific bacterial in seawater, or perhaps it is still there but in very small num- type in such diverse hosts and locations was that these bacteria bers. Yet another scenario is that today’s sponge-specific mi- became associated with an ancestral sponge prior to the evo- crobes were once a generalist marine species, thriving in all lution of current sponge classes (i.e., during the Precambrian). marine ecosystems, including sponges. Those strains that in- One should bear in mind, however, that the enormous com- habited sponges have since evolved to become genetically dis-

146 Microorganisms in marine sponges

tinct from their free-living counterparts. Support for these sequences from 21 sponges located around the world (Fig. 5). scenarios comes from another quarter, with various fatty acids We chose three of these sequences as an example, derived of likely microbial origin occurring in a wide range of sponges from the sponges Theonella conica (sampled from east Africa; irrespective of host phylogeny or geographic location (401, GenBank accession number AY701309), Aplysina aerophoba 403). The apparent absence of some of these biomarkers from (from the Mediterranean Sea; GenBank accession number marine sediments and seawater led to the suggestion that the AJ347056), and Antho chartacea (from southeastern Australia; compounds and their microbial producers have been present in GenBank accession number EF076240). The minimum pair- the sponges since ancient times (403). wise 16S rRNA similarity among these sequences (after cor- It is likely that any ancient sponge-microbe symbiosis would recting for different sequence lengths) is 97.9%. This is a very be obligate for one or both partners, potentially involving a minor difference when one considers the phylogenetically dis- reduction in microbial genome size if the symbiont has devel- parate hosts (the last two sponges are in different orders, while oped a nutritional dependence on its host. This has been dem- T. conica is in a different subclass) and their geographically onstrated for many obligate insect (e.g., see distinct locations. Even if one assumes that cyanobacteria references 252, 437, and 501), but it is unknown whether such evolve very slowly, we argue that greater sequence divergence tight host-symbiont coupling occurs in sponges. Integration of would be expected if these bacteria had indeed been living host and symbiont genomes was discussed in the sponge con- (separately) within their host sponges for 600 million years. text by Sara and colleagues (337), while a recent paper offers This should be especially true for endosymbiotic microorgan- evidence for lateral gene transfer from a fungus to the mito- isms, which are believed to evolve more rapidly due to in- chondrion of its host sponge (327). Such gene transfer would creased fixation of mutations within their small populations not be without precedent among marine invertebrates, as it is (259). These members of the “Ca. Synechococcus spongiarum” believed that the ascidian Ciona intestinalis laterally ac- cluster may therefore have a much more recent common ori- quired a cellulose synthase gene from a bacterium (253). gin, reflecting a role of horizontal (i.e., environmental) trans- Future genome sequencing of sponges and their microbial mission consistent with scenario 2 or 3 in Fig. 16. However, associates should offer valuable insights into the nature of consideration of other sequences within the same cluster can these symbioses. yield a quite different result. The two least similar sequences As noted earlier, not all sponge species harbor abundant within the “Ca. Synechococcus spongiarum” cluster are only microbial communities, and it is worthwhile to take a moment ϳ93% similar, suggesting a much older separation of these to consider these organisms. Freshwater sponges, for example, particular strains. A comparable degree of similarity is seen in typically contain a paucity of microbial associates, and it has comparing sequences from the “Ca. Synechococcus spongia- been suggested that this is due to an obligate requirement for rum” cluster with those from free-living relatives. We suggest sodium ions by the symbiotic bacteria (469). When freshwater that a combination of vertical and horizontal symbiont trans- sponges colonized their new habitat from the sea some 20 to 50 mission (scenario 2) could explain the observed data. Possible million years ago, it is presumed that existing symbionts were vectors responsible for horizontal symbiont transmission could lost. Many marine sponges also harbor only relatively small include sponge-feeding animals, such as fishes and turtles (205, numbers of microorganisms. These so-called low-microbial- 274), analogous to the coral-feeding fireworm Hermodice abundance sponges (148) often cooccur with the high-micro- carunculata, which acts as a vector for the coral pathogen Vibrio bial-abundance bacteriosponges, so habitat variation cannot be shilonii (386). invoked as an explanation for these differences. Whether these In our second example, we consider the “Poribacteria.” At sponges once contained, but later lost, large communities of first glance, there appears to be a strong case for an evolution- microbial symbionts is unknown. It is also unknown whether arily ancient relationship between these bacteria and their the (comparatively few) microorganisms in low-microbial- sponge hosts. The members of this monophyletic, exclusively abundance sponges are phylogenetically similar to those in sponge-specific bacterial lineage differ in their 16S rRNA se- their high-microbial-abundance counterparts. quences by up to 15% and are some 20% dissimilar to their Based on sequence information already at hand, the nearest nearest nonsponge relative (derived from Antarctic sediment) we can come to addressing these and the following hypotheses (Fig. 11). Such high divergence within the cluster, together is to consider estimated rates of 16S rRNA evolution for mem- with the low similarity to the next most similar known organ- bers of given sponge-specific clusters and to attempt to infer ism, is suggestive of an ancient symbiosis with sponges. How- when the last common ancestor of sponge-specific microbes ever, the two least similar “Poribacteria” sequences were taken from different sponges might have occurred. If one assumes from closely related (same family) sponges collected at the equal mutation rates in different bacterial lineages and asserts same Bahamas location, perhaps indicating horizontal symbi- thata1to2%16SrRNA sequence difference corresponds to ont transfer between these hosts. If the associations were an- approximately 50 million years of evolution (259), then se- cient and involved strict coevolution of host and symbiont, then quence differences of at least 10% would be required to place their respective phylogenies would be more congruent, with a common ancestor of these organisms back in the late Pre- the least similar microbes being hosted by the least similar (ϳ600 million years ago). Here we consider two sponges. Furthermore, the long naked branch leading to the examples, the cluster representing the cyanobacterium “Can- “Poribacteria” in the 16S rRNA tree could potentially be ex- didatus Synechococcus spongiarum” (426) and the “Poribacte- plained by faster rates of evolution in these bacteria. “Pori- ria” (100). The “Ca. Synechococcus spongiarum” cluster is one bacteria” are a sister phylum to the Planctomycetes (446), which of the largest of all sponge-specific sequence clusters, is well are sometimes believed to exhibit higher rates of evolution supported by all tree construction methods, and contains 52 than other lineages (392). Like the case for “Ca. Synechococ-

147 Part I

cus spongiarum,” a combination of vertical and horizontal sym- vertically transmitted, whereas secondary (facultative) symbi- biont transmission is thus the most likely scenario here, al- onts can be transferred either vertically or horizontally (329). though the acquisition of these bacteria exclusively from the Given that facultative symbionts can confer fitness advantages environment also cannot be ruled out. upon their hosts, maintenance of these populations—by what- Perhaps the most convincing evidence for a long-standing, ever mechanism—is of clear benefit. Interestingly, it was re- symbiotic relationship between sponges and at least some mi- cently shown that secondary symbionts in aphids can also be croorganisms comes from demonstrations of coevolution. De- transmitted via the sperm, yielding a different infection pattern spite difficulties in addressing this issue due to the phylogenetic from that which would be expected based on strictly maternal complexity of sponge-associated microbial communities, sev- transmission (237). As discussed in a subsequent section of this eral authors have now shown coevolution between sponges and article, both maternal and paternal transmissions of cyanobac- microbes. In the first study, the mitochondrial cytochrome terial symbionts have been documented for a marine sponge oxidase subunit 1 (CO1) gene and its bacterial homolog were (424). Another finding from the insect world which is relevant amplified from several halichondrid sponges and their associ- to our discussion is that certain bacteria can invade novel host ated bacteria (98). A CO1-based phylogenetic tree of six pu- species and form stable associations, perhaps using similar tatively alphaproteobacterial symbionts was largely congruent mechanisms for invasion to those found in with a tree containing sequences from the corresponding host (67). Provided that host chemical and immune defenses can be sponges, suggesting that cospeciation had occurred (although evaded, it therefore seems entirely plausible that marine mi- there also appeared to have been a host switch event at one crobes could invade, and establish themselves within, sponges point). Subsequent studies of the filamentous cyanobacterium from which they were previously absent. Oscillatoria spongeliae indicated a high degree of host specific- While phylogenetic trees of primary insect symbionts are ity for various dictyoceratid sponges, with evidence of cospe- congruent with those of their hosts, episodes of horizontal ciation as well as indications of some host switching (316, 396). transfer in secondary symbionts obscure the coevolutionary Ongoing studies of this system by Thacker and coworkers signal for these microorganisms. As mentioned in the preced- (R. W. Thacker, personal communication) should further elu- ing section, in which the cases of the “Poribacteria” and “Can- cidate the complex evolutionary relationships among these didatus Synechococcus spongiarum” were highlighted, the avail- tropical sponges and their cyanobacterial associates. Coevolu- able molecular evidence for sponge-associated microorganisms tion requires that the host and symbiont maintain close asso- supports a combination of vertical and horizontal transmis- ciations over evolutionary time, and as mentioned above, the sion (not just overall, but for specific individual lineages). mechanism by which this presumably occurs in sponges is ver- Another salient example is the alphaproteobacterial sponge tical transmission of microorganisms in host eggs or larvae. An associate represented in Fig. 14; this bacterium occurs additional point to consider at this stage is that the phylogeny widely in sponges (95, 453) and appears to be vertically of sponges themselves is not fully resolved (40). Molecular transmitted (95) yet is closely related to bacteria isolated data are often incongruent with traditional sponge taxonomy, from seawater. Issues of 16S rRNA sequence resolution which is based largely on morphological properties, such as notwithstanding (i.e., minor rRNA differences may hide ma- growth form and spicule characteristics (8, 37, 190). Accord- jor ecological or even genomic differences) (178, 323), this ingly, our understanding of symbiont evolution in sponges example underscores the complexities involved with consid- will continue to develop only in parallel with improvements erations of sponge-microbe evolution. in our knowledge of host phylogeny. A recently initiated CO1 Scenario 3: Environmental acquisition. In the third sce- sequencing project for taxonomically diverse sponges (www nario, putatively sponge-specific microorganisms are, in fact, .spongebarcoding.org) is a step in the right direction for also present in the surrounding seawater, but at such low abun- achieving the latter goal. dance that standard methods fail to detect them. Several mech- The final type of evidence for ancient, close associations anisms exist by which the same microbes may then be detected between sponges and microorganisms comes from the upon contact with a sponge. Firstly, it is possible that sponges record (43, 261, 377). Reef mounds constructed by siliceous absorb specific microbial types from seawater, a process which sponges and cyanobacterial mats, with the latter represented in would imply some degree of recognition of particular micro- part by still found today, flourished in (sub)trop- organisms (e.g., by the sponge’s innate immune system) (244). ical marine waters as far back as the early Cambrian (43). The Recognition of symbionts versus food bacteria has already fact that sponges and microbes closely coexisted hundreds of been proven experimentally (see the following section), and if millions of years ago is thus clear, but the nature of that a given microbe encounters favorable conditions (e.g., high interaction (e.g., whether microbes lived within sponge tissues) nutrients), it may multiply to the extent to which it can then be remains less certain. detected by the applied methods. Alternatively, a type of sub- Scenario 2: Parental and environmental symbiont transmis- tractive enrichment may occur, whereby those microbes which sion. Demonstrated vertical transmission is generally consid- cannot resist phagocytosis by sponge cells are consumed and ered a strong indicator of symbiosis, yet this does not rule out hardier bacteria (e.g., those with protective capsules) (482) the possibility of horizontal (e.g., environmental) transmission survive and are physically enriched near the choanocyte cham- of the same microbe as an additional mechanism. Indeed, this bers due to the sponge’s filtering activities. If such resistant phenomenon has already been shown for insect-bacterium bacteria are capable of out-competing other potential coloniz- symbioses (reviewed in reference 67), and here we borrow ers, they may establish themselves within the sponge tissue. from the well-developed literature on this topic. In aphids, the These possibilities can be placed under a banner of specific primary (obligate) bacterial symbiont Buchnera aphidicola is enrichment. Unspecific enrichment is another, at least theo-

148 Microorganisms in marine sponges

retical, alternative; in this case, microorganisms would simply sidered a sponge symbiont. Hunting for sponge-specific mi- be concentrated by sponges during filtering to the extent to crobes in other prolific filter feeders, such as ascidians (rather which they can then be detected by the applied methods. Al- understudied thus far, from a microbial perspective, but see though it is not easy to prove any of these hypotheses correct, references 239 and 413), may lead to the identification of these it is, in principle, even more difficult to prove them wrong. organisms from other, nonsponge sources. These arguments Finding a sponge-specific microbe actively living in the might seem to paint a grim picture for proponents of the ocean—independent from a host sponge—would lend support sponge-specific microbe concept, but the following must also to the enrichment hypothesis. The converse is less convincing: be considered. If it were really the case that such nonspecific if such cells are not detected outside sponges, it may simply enrichment of rare seawater microbes occurs in sponges, then reflect insufficient sampling. surely sponge-derived 16S rRNA gene libraries would be dom- Methodological considerations are of paramount impor- inated by other microbes which are known to be abundant in tance in discussing the environmental acquisition scenario. seawater. This does not happen. To illustrate the point, we use Even after many studies of marine microbiology, it still cannot again the example of the cyanobacterium “Candidatus Syn- be discounted that many or most of the so-called sponge- echococcus spongiarum”: if “Ca. Synechococcus spongiarum”- specific microbes are in fact also present in seawater, but only type organisms are very rare in the ocean but are concentrated at a low abundance which is not detected by standard methods. to sufficient levels to be detected in gene libraries from But how likely is it that sponge-specific microbes are actually sponges, then the exceedingly abundant (and closely related) present in other environments and that, due to methodological planktonic Prochlorococcus and Synechococcus strains should limitations, we simply fail to detect them? Given recent find- be much better represented. However, a cursory look at Fig. ings regarding the high level of diversity of uncommon micro- 5 reveals only a few such sequences from sponges. We thus organisms in the so-called rare biosphere (276, 376), the en- feel there is a strong case for rejection of the unspecific richment scenarios seem entirely plausible. Deep sequencing enrichment hypothesis. of seawater-derived 16S rRNA amplicons should provide fur- Regrettably, a lack of sufficient (appropriate) data prevents ther insights into the diversity of , per- us from ending this section with firm conclusions regarding the haps also yielding sequence types which are presently consid- origin of sponge-microbe associations. Even with almost 2,000 ered sponge specific. An interesting aside is that remarkably sponge-derived 16S rRNA sequences at our disposal, it is so- few of the Ͼ1,000 16S rRNA sequences obtained from the bering how little can actually be inferred about the evolution of Sargasso Sea metagenome (440) are closely related to those sponge-microbe associations. For almost every argument in sequences in sponge-specific clusters. That study employed a favor of an ancient symbiosis, there exists a rational counter- direct cloning approach and was thus free of PCR biases which argument which invokes (recent) environmental acquisition as might otherwise have resulted in the missing of certain se- the probable driver of the association. For example, a sponge- quence types. In this context, it will be very interesting to specific cluster at the end of a long naked branch in a phylo- examine the upcoming results of the Sorcerer II expedition genetic tree could reflect early divergence of this group from (www.venterinstitute.org), during which seawater samples are its relatives, or it may simply reflect insufficient sampling of being collected from around the world. Irrespective of what closely related lineages and/or accelerated evolutionary rates such studies find, if the sponge specificity of any microbe is to of the sponge-specific organisms. Additionally, while low levels be disproven, then it will be necessary to demonstrate activity of sequence divergence within a sponge-specific cluster argue of the said microbe outside a sponge host, as merely demon- against an ancient association, the corollary does not nec- strating its presence is not sufficient. essarily hold: extensive intracluster sequence divergence could Highly relevant to this discussion is an interesting point equally indicate a long, separate evolutionary history (e.g., made recently by R. Hill (154) regarding the abundance of among different hosts) or selective enrichment of diverse (but microorganisms in seawater and the potential consequences still monophyletic) organisms from seawater. So where do we for sponge microbiology. Central to this argument is the im- stand at the moment? At this stage, there is no clear indication mense filtering capacity of sponges, i.e., up to 24,000 liters of for scenario 1 (ancient symbioses), though this will be ad- seawater per day for a 1-kg sponge (443). Given that the typical dressed more satisfactorily in the future once additional infor- cell density of bacteria in seawater is about 106 cells/ml, then mation on sponge phylogeny is available. The existing data such a sponge could take in a staggering 2.4 ϫ 1013 bacterial point more towards scenario 2, whereby particular microbes cells per day. Thus, even if a specific bacterium is present may be passed vertically between generations, but with some in seawater at only 1 cell/ml (i.e., one-millionth of all cells horizontal exchange also occurring. Further consideration of present), then during a single day a sponge could still filter potential vectors for the latter process would be worthwhile. some 24 million cells of this organism from the Although we consider this scenario most likely, the supporting (154). The implications are obvious: an organism which is data are also consistent with scenario 3, i.e., environmental perennially extremely rare in seawater (and therefore never acquisition. This highlights a major current limitation from a detected by applied molecular methods) could conceivably be methodological perspective, namely, our inability to distin- concentrated within a pumping sponge (e.g., in the choanocyte guish between facultative sponge symbionts and specifically chambers, prior to phagocytotic ingestion) to an extent which enriched microorganisms. Presumably, the former prefer to is readily detectable by PCR or even hybridization-based meth- inhabit sponges but can tolerate conditions in seawater, while ods. If it occurs widely (but is undetected) in seawater, then enriched microbes may simply tolerate sponges long enough to one can imagine a situation in which it is easily detected from be detected by applied methods. Such apparently complex different sponges at different locations and erroneously con- patterns of microbial transmission are not without precedent in

149 Part I

the animal , with insect-bacterium symbioses provid- cytosis, followed by the release of lysozyme (400). Receptors ing a valuable framework for future considerations of sponge- for fungi (277) and viruses (466) also occur in sponges. For microbe symbioses. A well-studied marine system that has more detailed discussions of the various immune responses contributed greatly to our understanding of symbiont trans- and signal transduction pathways in sponges, the reader is mission—and, indeed, of host-microbe associations in gener- referred to recent reviews dealing with this topic (e.g., see al—is that of the squid Euprymna scolopes and its biolumines- references 244 and 246). cent symbiont, Vibrio fischeri (195, 230, 258, 442). V. fischeri is In another recent study from the Mu¨ller group, using S. acquired from the surrounding seawater by the juvenile squid, domuncula and its alphaproteobacterial symbiont SB2 as a which, via a remarkable stepwise process coined “winnowing,” model system, the importance of oxygenation of sponge tissue prevents all other bacteria from being established, culminating in mediating the relationship was demonstrated (241). Specif- in a monoculture of V. fischeri in its light organ (258). The ically, it was shown that strain SB2 grew preferentially on phylogenetically less complex microbial communities encoun- minimal media with the aromatic compound protocatechuate, tered in hosts such as insects and the squid light organ (typi- rather than glucose, as the carbon source. The bacterium can cally, in insects, there are one or two primary endosymbionts obtain protocatechuate in situ from the sponge, which pro- and one or two secondary endosymbionts) facilitate a depth of duces this and other diphenols via the activities of the enzyme understanding of these processes thus far not achievable for tyrosinase. Interestingly, tyrosinase activity and expression of the highly diverse microbial communities in sponges. the tyrosinase-encoding gene in S. domuncula, as well as the number of pcaDC genes in strain SB2 (responsible for bacterial utilization of protocatechuate and used here as a proxy for SB2 ECOLOGICAL ASPECTS: FROM SINGLE CELLS abundance on the surface [exopinacoderm] of the sponge), TO THE GLOBAL SCALE were all maximal under aerated conditions (241). Coupled with Establishment and Maintenance of the observed loss of SB2 cells from the sponge surface under Sponge-Microbe Associations low-oxygen conditions, it was asserted that the oxygen level is responsible for regulating the bacterial fauna in sponges. The mechanisms by which associations between sponges and Whether this type of mechanism is important in other sponge- microorganisms are established are not well understood. As microbe systems remains to be determined. discussed at length in the previous section, the fundamental The coexistence of microbial symbionts with bacterium-di- question of symbiont origin (i.e., whether symbionts were gesting archaeocytes in the sponge mesohyl has long interested passed down from an ancestral sponge or obtained contempo- sponge biologists. In a series of landmark experiments, Wilkin- raneously from seawater) remains unresolved, as do many of son and coworkers fed tritium-labeled sponge- and seawater- the mechanisms of sponge-microbe interactions and the regu- derived bacteria back to host sponges and found that sponge lation of microbial communities in these hosts. Extensive stud- symbionts passed through uneaten, whereas seawater bacteria ies by Mu¨ller and colleagues have attempted to address the were largely consumed (482). Two different mechanisms were underlying bases of sponge-microbe interactions at the molec- proposed to account for this: either (i) symbionts are specifi- ular level (36, 241, 243, 249, 399, 400, 444, 465). Sponges are cally recognized by the sponge and deliberately not ingested or often regarded as primitive animals, yet their morphological (ii) bacteria use extracellular masking capsules to avoid detec- simplicity belies the possession of a surprisingly complex im- tion by sponge cells (482, 483). While neither theory has been mune system (244). Indeed, the refinement of such a system tested explicitly (but see the preceding discussion on sponge has undoubtedly contributed to the success of sponges throughout immune responses), the latter explanation is in favor today, their long evolutionary history, especially when one consid- with several studies reporting the existence of slime layers and ers the immense numbers of (potentially pathogenic) microor- sheaths on symbiotic bacteria (106, 427, 473). The results of ganisms to which they are exposed due to their filter-feeding recent experiments by the Hentschel group were consistent activities. Detailed studies of the Adriatic sponge Suberites with earlier findings: seawater-derived bacteria were consumed domuncula have revealed immune responses against both by Aplysina aerophoba some 2 orders of magnitude faster than gram-negative (36, 243, 464) and gram-positive (400) bacteria, was a consortium of sponge-derived bacteria (459). In addi- illuminating one means by which sponges may select for and tion, when a green fluorescent protein-labeled food bacterium against certain microbes from the surrounding environment. In was fed to the sponge, it was rapidly digested within the sponge the case of the former, exposure of S. domuncula to the bac- tissues. All of these findings carry interesting implications for terial endotoxin lipopolysaccharide (LPS)—derived from the evolution of sponge-microbe associations (also see the gram-negative cell walls—elicited an increase in synthesis of previous section). If presumed symbionts are not taken from two compounds with pronounced antibacterial activity (243). the seawater (i.e., either as colonizers or as a food source), Confirmation that the compounds were indeed synthesized by then this suggests vertical transmission as the mechanism by the sponge was obtained by cloning of the gene encoding a key which these associations are maintained. enzyme in the relevant biosynthetic pathway. A receptor for It is now established beyond doubt that many sponges (at LPS on the sponge cell surface was later identified, as was a least among those in marine environments) harbor diverse and signal transduction pathway which is upregulated upon expo- abundant microbial communities. What is far from established sure to increased LPS levels (464). The immune response of S. is how, if at all, the composition and density of these commu- domuncula to gram-positive bacteria takes a quite different nities are regulated. The potential role of phages and protozoa form: upon exposure to peptidoglycan in the bacterial outer in regulating microbial communities within sponges is of inter- , the sponge responds with a rapid activation of endo- est, but virtually nothing is known about this to date (but see

150 Microorganisms in marine sponges

reference 211). Predatory bacteria, perhaps related to the deltaproteobacterial genus Bdellovibrio, could also be involved in structuring microbial communities in sponges (468). At least for cyanobacteria, it has been suggested that their abundance is directly proportional to the number of sponge cells, implying some degree of influence by the sponge over symbiont growth and reproduction (477). The high photosynthetic rates of cya- nobacteria in sponges (see also the following section) should, with all things being equal, result in cyanobacterial growth to the extent that host tissues would be overwhelmed. It is thus likely that the sponge exerts some control over its symbiont populations, with several mechanisms being proposed, includ- ing the following: sponges consume excess symbionts, sponges eject symbionts when stressed, the host sponge steals photo- synthate from the cyanobacteria, and sponges starve the sym- bionts (477). With some debate over the extent of sponge consumption of symbionts and no evidence for expulsion of excess symbionts, Wilkinson argued that the last two scenarios (steal and starve) are most likely. There is strong evidence for stealing of photosynthate from symbionts in other systems (e.g., coral-zooxanthella [128] and freshwater Hydra-Chlorella [232] symbioses), and it seems plausible that sponges may also FIG. 17. Vertical transmission of microbial symbionts by a marine produce some sort of host release factor to induce the release sponge. A transmission electron micrograph of a Chondrilla australien- of large quantities of fixed carbon from the cyanobacteria. sis larva is shown, indicating a diverse range of bacterial morphotypes. Along these lines, a host release factor was recently described Bar ϭ 1 ␮m. (Modified from reference 420 with permission of the for the symbiosis between the sponge Haliclona cymaeformis publisher.) and its macroalgal symbiont Ceratodictyon spongiosum (129). In contrast, the starve hypothesis holds that if a sponge can somehow restrict the symbiont’s access to essential nutrients, two ways: (i) microbes can be recruited from the surrounding such as nitrogen and , then symbiont protein syn- water by filter feeding (i.e., horizontal or environmental trans- thesis and cell division would be restricted. Consequently, an mission) or (ii) microbial symbionts can be passed on from the excess supply of carbon-rich photosynthate would be excreted parent sponge via reproductive stages (i.e., vertical transmis- from the symbiont (156, 477). To the best of our knowledge, sion) (Fig. 17). Although horizontal transmission of symbionts neither scenario has been proven unequivocally or disproven has been demonstrated convincingly for several marine symbi- for any sponge-microorganism association. More generally, re- oses (e.g., squid-Vibrio fischeri [258] and markably little is known about communication, or chemical tubeworm-chemoautotrophic bacterium [257] symbioses), it is cross talk, between sponges and their microbial associates. the latter mechanism which has received most attention for Marine sponges produce a wide variety of secondary metabo- sponges. Bacteria have now been found in embryos or larvae lites, some of which could potentially enable them to select for from all three classes of Porifera (see reference 97 and refer- or against particular types of microorganisms (185) (although ences therein), including species with highly varied reproduc- the nonspecific nature of many antimicrobial compounds sug- tive strategies. Sexual reproduction in sponges involves either gests that such selection may generally be limited to broader vivipary (where larvae are brooded within the animal) or classes of microbes, e.g., gram-positive versus gram-negative ovipary (whereby eggs, generally fertilized externally, develop bacteria). Conserved bacterial signaling systems, as exempli- outside the sponge). Evidence for vertical transmission of bac- fied by the acyl homoserine lactone (AHL) regulatory systems teria has been reported for both types (97, 116, 118, 181, 362, of many gram-negative bacteria (111, 369), often mediate col- 419, 422, 431), while , i.e., budding, could onization-related traits (e.g., biofilm formation, swarming, and also contribute to symbiont transfer in some species (146). virulence) and offer one means by which sponges could interact Indeed, gemmules, the asexual buds of freshwater sponges, or interfere with bacteria. Bacteria capable of AHL production contain symbiotic zoochlorellae in at least some species (372), have already been reported from marine sponges (361, 389), as while a bud protruding from the surface of the marine sponge have other putative signaling molecules, such as diketopipera- Tethya orphei contained a symbiotic cyanobacterium (117). zines (1, 162, 182). It is highly likely that sponges produce The vast majority of reports dealing with vertical transmis- metabolites which allow them to disrupt AHL-regulated phe- sion in sponges have been based on transmission electron mi- notypes, as shown for the macroalga Delisea pulchra (126, 127, croscopy (TEM) observations. Such studies have contributed 223, 224) and various terrestrial plants (302, 393). Indeed, greatly to our understanding of this phenomenon, with the inhibition of bacterial swarming by chemical extracts from identification of several mechanisms by which symbiotic mi- sponges has recently been shown, though it has yet to be crobes can be incorporated from maternal mesohyl tissue into clarified whether this is an AHL-specific effect (184). eggs or embryos (reviewed in reference 97). These include Associations between sponges and microorganisms can be phagocytosis of microbial cells by the oocyte directly from the maintained over different generations in either of the following adult mesohyl as well as transfer of microbes from parent

151 Part I

sponge to embryo along an “umbilical cord.” Intriguingly, in improve our understanding of the mechanistic bases of these the Australian sponge Chondrilla australiensis, eggs containing symbioses. a cyanobacterial symbiont (of the “Candidatus Synechococcus Whatever the underlying mechanisms for the establishment spongiarum” type [426]) were distributed throughout the and maintenance of sponge-microbe associations, it is appar- sponge mesohyl, whereas cyanobacteria are normally confined ent that in many cases such associations are highly stable and to the better-illuminated periphery, or cortex, of the sponge resistant to external disturbance (reviewed in reference 148). (422). Nurse cells, probably derived from choanocytes (425), In at least some other instances (e.g., see reference 457), this is have been invoked as a possible mechanism by which cya- not the case. Briefly, neither starvation, exposure to antibiotics, nobacteria are transported to the eggs (422, 424). These cells, nor transplantation to different depths could elicit major which fuse with eggs and release their contents (including changes in bacterial community composition in Aplysina aero- cyanobacteria) into the egg cytoplasm prior to spawning, pre- phoba (105) and Aplysina cavernicola (407). Similarly, even sumably phagocytose the symbionts in the cortex before mov- translocation of Aplysina fistularis from its natural depth of 4 m ing to the developing eggs deeper within the sponge. Remark- to a new depth of 100 m was not enough to significantly affect ably, Usher and coworkers were also able to demonstrate the cyanobacterial abundance in this sponge (although the sponge presence of cyanobacteria in sperm cells, indicating that both did die at depths of Ͼ100 m) (222). In contrast, this change parents are capable of transferring symbionts to offspring resulted in a loss of cyanobacterial symbionts from the cooc- (424). Sponges of the genus Chondrilla were also the subject of curring sponge Ircinia felix. The importance of cyanobacterial another recent TEM study (which additionally employed im- symbionts—at least to some sponges—was recently demon- munocytochemical techniques), in which vertical transmission strated by Thacker in a series of elegant field experiments of an endosymbiotic yeast was shown (221). (394). To test the hypothesis that a greater benefit to the host The drawback of the TEM approach is that, with some is derived from more specialized symbionts, shading experi- exceptions (e.g., cyanobacteria [423, 424]), even phylum-level ments were conducted with the tropical sponges Lamellodys- idea chlorea Os- identification of the relevant microorganisms is not possible (containing the host-specific cyanobacterium cillatoria spongeliae) and Xestospongia exigua (which contains due to an insufficient number of distinguishing morphological the generalist symbiont “Ca. Synechococcus spongiarum”). characters. Multiple bacterial morphotypes have been ob- Shaded individuals of the encrusting sponge L. chlorea lost served in sponge larvae (suggesting transmittance of a complex Ͼ40% of their initial area, but the a concentration assemblage) (e.g., see reference 424), yet little or nothing is in the remaining sponge tissue (a proxy of cyanobacterial abun- known of their phylogenetic affiliations. The recent application dance) did not change, implying a close, putatively mutualistic of molecular techniques in this area (95, 266) thus offers the relationship between the sponge and O. spongeliae. In contrast, potential for exciting new insights into the phylogenetic (and, X. exigua lost relatively little mass but did lose many of its “Ca. in principle, metabolic) complexity of transmitted microbial Synechococcus spongiarum” symbionts, suggesting that this re- assemblages. A 16S rRNA gene library constructed using cya- lationship is less tight-knit than the other (394). While the nobacterium-specific PCR primers confirmed the presence (as mechanism by which symbionts are lost from X. exigua is un- indicated by TEM) of a single cyanobacterial type in both clear, the existing data do suggest that the specialist O. spon- Diacarnus erythraenus larvae and adults of the Red Sea sponge geliae provides a greater benefit for its host sponge than does (266). The transmitted cyanobacterium is highly similar to the the generalist “Ca. Synechococcus spongiarum” for its host. aforementioned “Ca. Synechococcus spongiarum”-type symbi- Although the necessary data are currently lacking, one could ont of Chondrilla australiensis. A range of molecular tech- speculate that the degree of host specificity (of individual sym- niques were used to examine the bacterial community in larvae bionts or even entire communities) may explain some of the of the Caribbean sponge Mycale laxissima, revealing a much different results obtained from previous perturbation experi- more diverse population than that recovered by cultivation ments (148). More generally, the extent of host specificity efforts (95). A single alphaproteobacterium, related to among marine -associated microbes may have sub- Pseudovibrio denitrificans and previously reported from many stantial implications for microbial diversity on a wider scale sponges (Fig. 14), was the only bacterium from the larvae that (390). If most symbionts are highly host specific, then their could be grown on a standard marine medium. In contrast, overall diversity will be much higher than if the same hosts sequences representing a diverse assemblage comprising Acti- harbor mostly generalist species. Finally, Roberts and cowork- nobacteria, Bacteroidetes, Cyanobacteria, Planctomycetes, and ers recently reported the results of experimental manipulations Proteobacteria (including the isolated alphaproteobacterium) with the photosynthetic symbiont-containing Australian sponge were recovered from a 16S rRNA gene library based on DNAs Cymbastela concentrica (322). Shade and, to a lesser extent, isolated from the larvae (95). Similarly diverse microbial com- treatments (both designed to mimic the physical effects of munities were found in larvae of the sponges Corticium sp. sewage effluent discharge) led to lowered chlorophyll a con- (366) and Ircinia felix (352), using 16S rRNA-based ap- centrations within the sponge, while increased salinity and nu- proaches, such as gene libraries, FISH, and DGGE. Broad trient loads had negligible effects. congruence between larva- and adult-associated microbial communities in these studies indicated that a significant subset Physiology of Sponge-Associated Microorganisms of the resident microbes is transferred in this way. Future molecular studies could provide information on the metabolic A lack of pure cultures for most sponge-associated micro- properties of the transferred symbionts (e.g., via analysis of organisms has contributed to a paucity of knowledge about functional genes or FISH-microautoradiography), which should their physiological characteristics. What we do know has arisen

152 Microorganisms in marine sponges

from a combination of the existing pure-culture studies, pro- thetic symbionts to host metabolism (324). Conversely, in at cess data, and inferred metabolic properties from analysis of least one case, it appears that diatoms in Antarctica may par- 16S rRNA, functional genes, and metagenome data. Collec- asitize the sponge host, using its metabolic products as an tively, microbes in sponges are capable of, among other pro- energy source (16). The sponge Cymbastela concentrica in cesses, photosynthesis, methane oxidation, nitrification, nitro- southeastern Australia contains high densities of diatom-like gen fixation, sulfate reduction, and dehalogenation. Here we cells in the illuminated periphery (M. W. Taylor, unpublished summarize the current knowledge by first examining, in turn, data), but further work is required to elucidate whether this the major nutrient cycles within sponges. association is phototrophic in nature. The occurrence in this Carbon. Heterotrophy is a common form of carbon metab- sponge of at least some cyanobacteria in addition to the dia- olism in sponges, either via consumption of microbes from toms (Longford et al., unpublished data) will complicate efforts seawater or via microbial uptake of dissolved organic carbon in this direction. Some freshwater sponges (e.g., Spongilla (495). However, for many sponges, particularly those in trop- lacustris) contain zoochlorellae, and although the symbiosis is ical regions, carbon metabolism centers around the activities of not obligate (aposymbiotic individuals occur in areas of deep photosynthetic microorganisms, such as cyanobacteria (11, 58, shade), it appears that algal photosynthesis can contribute to 59, 381, 470, 474, 477, 486). Many tropical sponges contain host metabolism and growth (109, 333, 475). substantial populations of these oxygenic , and no- An unusual form of nutritional symbiosis is that between where is the contribution of microbial symbionts to the host methanotrophic bacteria and deep-sea cladorhizid sponges sponge more evident than in this case (see also the next sec- (428, 429, 431). These remarkable sponges, which possess no tion). Translocation of photosynthates (mostly as glycerol) aquiferous system but instead prey on tiny swimming organ- from cyanobacteria to the host has been shown for marine isms, are believed to obtain a significant portion of their nu- sponges (476), while glucose produced by a chlorella-like green trition from the consumption of methanotrophs. Methane alga was passed to its freshwater sponge host, Ephydatia flu- serves as a carbon source and substrate for energy production viatilis (475). Phototrophic sponges—those whose carbon nu- in methanotrophs, and in this particular system it is derived trition depends heavily on cyanobacterial symbionts—receive from a deep-sea mud volcano (429). In other sponges, the Ͼ50% of their energy requirements from cyanobacteria (474), presence of methanogenic archaea may lead to methane pro- allowing these species to thrive in the low-nutrient, high-light duction within anoxic zones. A 16S rRNA gene sequence af- areas commonly found on tropical reefs. On the Great Barrier filiated with the methanogens of the phylum Euryarchaeota Reef, phototrophic sponges comprise approximately half of (456) is the sole piece of evidence for this at present, but the the total sponge biomass on outer reefs, where the water is documented existence of anoxic microhabitats within sponges cleaner, but are much less common inshore, where terrestrial (159) suggests that these associations could be more wide- runoff and turbidity are greater (470, 485). Similarly, phototro- spread. Other chemoautotrophic microbial processes that have phic sponges are largely absent from Caribbean reefs, where been observed in sponges and may also contribute to sponge only small numbers of sponge-associated cyanobacteria are nutrition are nitrification and sulfur oxidation. These are dealt present (470). Phototrophy has also been demonstrated for at with in the following sections. least one temperate sponge (57), while numerous others are Nitrogen. After carbon, nitrogen is the most important nu- known to contain photosynthetic symbionts (321, 336, 390, trient for life, as it is required for the synthesis of amino acids 430). The sponge Cymbastela sp. from temperate South Aus- and, subsequently, proteins. In oligotrophic waters where ni- tralia was capable of compensating photosynthetically (i.e., the trogen levels are low (e.g., coral reefs), symbiotic microorgan- rate of photosynthesis equals its rate of respiration) at a 4- to isms may contribute to the nitrogen budget of sponges via

5-m depth in winter, while it was a net producer at the same fixation of atmospheric nitrogen, N2 (479, 484). The first evi- depth in summer (57). In contrast, Great Barrier Reef sponges dence for this came from measurements of nitrogenase activity may derive much of their nutrition from photosynthetic sym- in three Red Sea sponges (479). The activity of this enzyme, bionts as deep as 15 to 30 m, due to the clearer water and, the catalyst for microbial N fixation, was estimated using an therefore, decreased light attenuation (58). Some sponges are acetylene reduction test (for caveats, see reference 484) and apparently obligate phototrophs, with their lower depth limits could be measured only in Siphonochalina tabernacula and determined by the availability of light for photosynthesis (58). Theonella swinhoei, both of which contained cyanobacteria. In Others function as mixotrophs, combining symbiont-derived contrast, Inodes erecta, which contained only noncyanobacte- nutrition with filter feeding, while still others contain no pho- rial microorganisms, showed no evidence of N fixation. Addi- toautotrophic symbionts and derive all of their carbon nutri- tionally, nitrogenase activity was higher in illuminated tissue tion from filter feeding (477). than in that maintained in the dark and did not correlate with The extent to which other photosynthetic associates (e.g., the abundance of the heterotrophic bacterial communities in S. diatoms, dinoflagellates, and phototrophic sulfur bacteria) tabernacula and T. swinhoei. Taken together, these data sug- contribute to carbon cycling within sponges is less clear. The gested that nitrogenase activity was due mainly to the presence Mediterranean sponges Cliona viridis and Cliona nigricans both of cyanobacteria, many of which are capable of N fixation contain symbiotic dinoflagellates (zooxanthellae), and for C. (479). A subsequent study provided more concrete proof of N viridis, at least, it appears that sponge metabolism depends on fixation in sponges by demonstrating incorporation of the sta- 15 the photosynthetic activity of these symbionts (353). Indeed, ble isotope N2 into various amino acids in Callyspongia muri- the growth of C. viridis was greater in individuals maintained cina (484). Whether microbial N fixation is of major ecological under natural light conditions than in those maintained in significance for sponges remains uncertain, but it does appear constant darkness, reflecting the contribution of photosyn- that its occurrence in sponges is not limited to cyanobacteria.

153 Part I

FIG. 18. Current state of knowledge about the nitrogen cycle in sponges. Thick arrows signify those processes which have been demonstrated in sponges; references (given in parentheses) pertain to either the process or the implicated microorganisms. PON, particulate organic nitrogen.

Heterotrophic nitrogen-fixing bacteria were reported from a [193, 300]). Nitrite oxidizers belonging to the Nitrospira genus Halichondria sp. (367), and the nifH gene (encoding a subunit are frequently recovered in 16S rRNA gene surveys of sponges, of the nitrogenase reductase enzyme) has been amplified from yet in at least one case (80), extensive release of nitrite indi- both alpha- and gammaproteobacteria inhabiting several Ca- cated that oxidation of ammonia and nitrite could be uncou- ribbean sponges (N. M. Mohamed, Y. Tal, and R. T. Hill, pled. Interestingly, ammonia-oxidizing archaea, whose exis- presented at the 11th International Symposium on Microbial tence was just proven in 2005 (192), also exist in sponges. Ecology, Vienna, Austria, 20 to 25 August 2006). Metagenomic reconstruction of “Candidatus Cenarchaeum Nitrification in sponges has also received attention. The two symbiosum,” the abundant and yet uncultivated crenarchaeote steps of nitrification, i.e., the biological conversion of ammonia in the Californian sponge Axinella mexicana (294), revealed the to nitrite and then to nitrate, are catalyzed, in turn, by ammo- existence of ammonia monooxygenase (amoA) genes (required nia-oxidizing and nitrite-oxidizing microorganisms (33, 194). for ammonia oxidation) in this organism (135), while PCR- Ammonia, which can be toxic to eukaryotes, is a metabolic based surveys for amoA indicated that archaeal ammonia ox- waste product and could accumulate within sponge tissues, idizers are widespread in marine sponges (D. Steger et al., particularly during periods of low pumping activity. The re- unpublished data). Whether archaea or bacteria are the key lease of nitrate (and in some cases nitrite) from incubated ammonia oxidizers in sponges remains to be determined, but in sponges provided the first indication of nitrification within at least one system (soils), ammonia-oxidizing archaea appear these organisms, with estimated rates often far exceeding those to greatly outnumber their bacterial counterparts (204). for other benthic substrata (63, 80). These results suggested Numerous gaps remain in our knowledge of nitrogen cycling the presence of nitrifying microorganisms, and indeed, 16S in sponges (Fig. 18). For example, the existence of anoxic rRNA sequences from both ammonia-oxidizing betaproteo- zones in at least some sponges (159) suggests the potential for bacteria (79; Taylor et al., unpublished data) and nitrite- both denitrification and anaerobic ammonium oxidation (anam- oxidizing bacteria of the genus Nitrospira (Fig. 10) (146; Long- ford et al., unpublished data) have been recovered in molecular mox). Neither process has been reported for sponges thus far, surveys of sponges. The widespread presence of Nitrospira in and our own efforts to amplify 16S rRNA genes from known sponges may indicate low nitrite availability in these hosts, as anammox bacteria from several sponges have yielded no re- members of the Nitrospira typically favor low-nitrite habitats sults. Denitrification is catalyzed by a phylogenetically diverse (356, 448). Nitrifying microorganisms are among the few for range of microorganisms, and it is risky to infer the ability to which metabolic capabilities can generally be inferred from denitrify from 16S rRNA sequence data alone. Nonetheless, it 16S rRNA data, and sequences representing several types of is worth noting that a common alphaproteobacterial associate ammonia-oxidizing bacteria–in the genera Nitrosospira and Ni- of marine sponges (95, 147, 453) (Fig. 14) is very closely related trosomonas–-were identified from the Australian sponge Cym- to the marine denitrifier Pseudovibrio denitrificans (368), and at bastela concentrica (Taylor et al., unpublished data). The find- least some of the sponge-derived strains have also tested pos- ing of only eutropha/europaea-affiliated ammonia itive for denitrification (95). The role of sponge filter feeding in oxidizers in a previous study of six tropical sponges (79) may providing particulate organic nitrogen is also of interest (312), have been due to the use of overly specific PCR primers (both with evidence that uptake of ultraplankton can yield sufficient of the primers used have mismatches to all Nitrosospira-affili- nitrogen to sustain both the tropical sponge Haliclona cymae- ated ammonia oxidizers and also to many Nitrosomonas spp. formis and its macroalgal symbiont (72, 288).

154 Microorganisms in marine sponges

Sulfur. Several lines of evidence point to the widespread but these sequences do not appear to have been deposited in occurrence of sulfur-metabolizing microorganisms in sponges. GenBank and were therefore not included in our analyses. For starters, two of the key microbial players in the sulfur cycle, Analysis of functional genes [e.g., dsrAB, encoding the dissim- namely, sulfate reducers and sulfur oxidizers, have been found ilatory (bi)sulfite reductases] (447, 502) would be one way to in multiple sponges. Sulfur-oxidizing bacteria from the families gain further insights into the composition of sulfur-metaboliz- Rhodospirillaceae and Chromatiaceae (Alpha- and Gammapro- ing microbial guilds within sponges. teobacteria, respectively) were isolated from Ircinia sp. and A final aspect of sulfur metabolism that has received much Euspongia officinalis in the 1970s (173). In that paper, the attention in marine systems is that of dimethylsulfoniopropi- bacteria were referred to as phototrophic sulfur bacteria, and onate (DMSP) and its cleavage product, dimethylsulfide (DMS). additional, earlier isolations of green sulfur bacteria (phylum These compounds are thought to play a role in global climate Chlorobi) were also discussed (see reference 173 and the Eim- regulation (497), while DMSP may also protect marine algae hjellen [1967] citation within). FISH signals for Chlorobi were and invertebrates from herbivores/predators and oxidative later found in the Great Barrier Reef sponge Rhopaloeides damage (384, 436). In a recent survey of DMSP content in a odorabile (458). The above-mentioned sulfur bacteria oxidize variety of coral reef invertebrates, high levels in corals were reduced sulfur compounds such as hydrogen sulfide. This sub- attributed to symbiotic zooxanthellae, while the much lower strate is presumably derived from the activities of sulfate-re- DMSP concentrations typical of sponges were presumed to be ducing bacteria, which have also been obtained from sponges diet derived (435). It is currently unknown whether the levels (159, 160, 173, 225, 358). An endosymbiotic sulfur cycle com- in sponges are sufficient to play a role in predator deterrence prising sulfate-reducing and sulfide-oxidizing bacteria has al- or whether those sponges with symbiotic zooxanthellae have ready been demonstrated for a marine oligochaete (83), and higher DMSP contents. the above data suggest that a similar process takes place in at Other aspects of microbial metabolism in sponges. In con- least some sponges. trast to the case for several major chemical elements (namely, The most extensive work on sulfur metabolism within carbon, nitrogen, and sulfur), to our knowledge virtually noth- sponges has been conducted by Hoffmann, Reitner, and col- ing is known about phosphorus cycling within marine sponges. leagues (159, 160, 225, 310, 358). They detected sulfate-reduc- We assume that sufficient phosphorus is obtained from the ing bacteria by FISH in the Mediterranean sponges Chondrosia sponge’s diet of microorganisms. reniformis and Petrosia ficiformis (225, 358), as well as in the The degradation of halogenated chemicals within the Med- cold-water sponge Geodia barretti (159, 160). In G. barretti, iterranean sponge Aplysina aerophoba was the subject of an FISH detection of sulfate reducers belonging to members of interesting recent study (3). This and other sponges are rich the genera Desulfoarculus, Desulfomonile, and/or Syntrophus sources of brominated compounds such as bromophenols and (estimated abundance, 1.8% of the total bacterial community) bromoindoles, and it was predicted that microorganisms within was complemented by isotopic measurements of sulfate reduc- such sponges may be capable of dehalogenation. Indeed, by tion rates and analysis of oxygen profiles within the sponge establishing enrichment cultures from sponge tissue, in the (159). Sulfate reduction is an anaerobic process, and through presence or absence of various electron acceptors, the authors the use of microelectrodes (354), these authors were able to of that study were able to demonstrate reductive debromina- demonstrate the presence of anoxic zones within the sponge, tion under methanogenic and sulfidogenic, but not denitrify- particularly during periods of pumping inactivity (159). The ing, conditions (3). Antibiotic inhibition of dehalogenation ac- estimated sulfate reduction rates in G. barretti, of up to 1,200 tivity indicated that it was the microbes, not the sponge, which 2Ϫ 3 Ϫ1 nmol SO4 cm sponge day , are among the highest re- were responsible. corded in natural systems. Intriguingly, analysis of lipid bio- The production of a wide range of secondary metabolites by markers suggested that bacterially derived carboxylic acids sponge-associated microorganisms is well known. We provide (perhaps from sulfate reducers) may be transferred to the host examples of some pharmacologically relevant metabolites in a for subsequent synthesis into other compounds (159). The later section (see “Biologically Active Chemicals from Marine accumulation of toxic hydrogen sulfide was also addressed by Sponge-Microbe Consortia and Their Commercial-Scale Sup- Hoffmann and coworkers, who calculated that the activities of ply”) and, immediately below, outline the potential benefit(s) sulfide-oxidizing bacteria could, together with chemical reoxi- of these metabolites to the sponge-microbe association. dation processes and the use of oxidized iron from seawater as an electron acceptor, be sufficient to balance microbial sulfide The Varied Nature of Sponge-Microbe Interactions production. Although it is pure speculation at this stage, it is also possible that sulfur-oxidizing symbionts enable sponges to Sponges and the microorganisms living within and around occupy sulfide-rich environments. The base of the Micronesian them display the full gamut of interactions, from microbial sponge Oceanapia sp., for example, can be buried up to 20 cm pathogenesis and parasitism (sometimes resulting in sponge deep in the sediment (360), where anoxic conditions with high death) to microbes as the major food source for heterotrophic sulfide concentrations may prevail. sponges and to mutualistic (or at the very least commensalistic) Interestingly, 16S rRNA gene sequences which are highly associations in which both partners appear to benefit. We first similar to those from known sulfate reducers (e.g., Desulfobac- consider the putative benefits of symbiotic microorganisms to terium or Desulfomicrobium spp.) (Fig. 8) have only rarely been the host sponge. recovered from sponges. Ahn and colleagues did find Desulfo- Mutualism/commensalism. It is clear that sponges benefit vibrio-related organisms in enrichment cultures grown on greatly from the diverse metabolic properties of their associ- Aplysina aerophoba-derived brominated phenolic compounds (3), ated microorganisms (see the preceding section). The provi-

155 Part I

sion of photosynthates and (perhaps to a lesser extent) fixed strated in laboratory experiments (179), it is less clear whether nitrogen from cyanobacteria (11, 474, 476, 477, 479, 486) is ammonia or nitrite ever accumulates to sponge-harming levels presumably a key factor in the ecological success of many in nature. Microelectrode studies to address such questions sponges on nutrient-poor tropical reefs. Cyanobacterial sym- (354) would be of great interest and should further our under- bionts may be equally important to juvenile and adult sponges. standing of the role of nitrifying microorganisms in sponges. Sponge larvae are generally thought to be lecithotrophic (i.e., Also of interest will be the data derived in the future from nourished from finite stored nutrients) (219), with no capacity sponge genome projects. These should help to identify possible for filter feeding (though some may assimilate dissolved or- absent metabolic pathways in the host, whose functions may ganic matter from seawater [175]), so the energy gained from instead be filled by symbiont-derived factors. photosynthetic cyanobacteria should contribute to (i) Further putative benefits for sponges from their microbial and larval longevity in the water column (424) and (ii) (once partners include increased structural rigidity (due to mucous sponges are settled) the rapid growth required to outcompete production by bacteria) (472), direct incorporation of dissolved algae and other photosynthetic organisms for substratum in organic matter from seawater (480, 495), digestion and recy- illuminated areas (477, 478). Larval mortality may conse- cling of insoluble sponge collagen (481), and microbial pro- quently be lower for those harboring cyanobacterial symbionts. duction of secondary metabolites that are of use to the host. In The importance of photosynthetic symbionts to their hosts is several cases, production of bioactive metabolites has tenta- evident in the typically flattened morphologies of phototrophic tively been ascribed to bacterial symbionts, and these may sponges, with the thinner species containing dense accumula- serve to protect the sponge from pathogens, predators, and tions of cyanobacteria throughout the tissue. In contrast, mixo- foulers (e.g., see references 147, 351, and 417). In some other trophic sponges—those which utilize both filter feeding and cases, molecules produced by microbial symbionts could po- photosynthetic symbionts for nutrition—may reduce their re- tentially be used as precursors for the of defense liance on symbionts with age. Juveniles possess a high propor- metabolites by sponges. Whatever the exact mechanism, it is tion of symbiont-containing tissue, which reduces as the likely that the chemical defenses of many sponges include both sponge grows thicker and increases the amount of filter-feed- host- and symbiont-produced metabolites (see also “Harming ing tissue (474, 478). Interestingly, photosynthe- the host: pathogenesis, parasitism, and fouling” and Bio- sis can also bring with it certain costs for the sponge, such as technology of Sponge-Microbe Associations: Potential and the following: (i) morphological adaptation for improved light Limitations). capture may occur at the expense of filter-feeding capacity The observant reader may have noticed that the preceding (477), and (ii) oxidative stress may result from the presence of discussion deals almost exclusively with putative benefits for high levels of photosynthetically produced molecular oxygen, the host, with little mention of advantages for the symbiont(s). necessitating an enhancement of antioxidant defenses com- Even when the benefits to the host sponge are obvious (e.g., in pared with those of asymbiotic specimens (303, 304). Another phototrophic sponges), it is not necessarily clear what benefit role for cyanobacteria and their has also been pro- the microbial partner derives from the association. It is thus posed, namely, protection of sponges from excessive illumina- difficult, and often impossible, to confidently assign a mutual- tion (336). Although this role has not been proven experimen- istic rather than just a commensalistic label to a sponge-mi- tally, one expects that this could be particularly important for crobe association. Presumed benefits for microbial symbionts intertidal species, where radiation (UV and photosynthetically of sponges include a generous supply of nutrients, as well as active radiation) is especially high (381). The documented shelter—from predators or high light levels—in the sponge occurrence of UV-absorbing mycosporine-like amino acids tissue (336). in sponges harboring cyanobacteria (e.g., Dysidea herbacea Microorganisms as a food source for sponges. With the [13]) also supports the hypothesis of shading by the symbionts. primary exception of phototrophic sponges (described above), Microbial metabolism may benefit the host sponge in other most sponges are thought to obtain the bulk of their carbon ways. As mentioned earlier, Hoffmann and colleagues (159) nutrition via the consumption of microorganisms from the described the likely transfer of carboxylic acids from anaerobic water column (though uptake of dissolved organic carbon may bacteria to the sponge Geodia barretti. Methanotrophic bacte- also be significant [e.g., see reference 495]). Bacteria (including ria may supplement the nutrition of non-filter-feeding, carniv- both cyanobacteria and presumed ) as well as orous sponges in methane-rich deep-sea habitats (429, 431), eukaryotic microalgae can satisfy the entire food requirements while symbiotic zooxanthellae (dinoflagellates) enhance boring of sponges (307), with the potential for dense sponge commu- and growth rates in clionid sponges (153). Elimination of toxic nities to significantly deplete the surrounding water of micro- metabolic by-products is another possible role played by sponge- bial cells (289, 309). Early studies of particle feeding in sponges associated microbes. The sulfur-oxidizing bacteria mentioned indicated that as much as 96% of bacterial cells were removed above oxidize reduced sulfur compounds, such as highly toxic from the inhalant seawater by the filtering activities of the hydrogen sulfide, to less harmful forms. Sulfide may accumu- sponge (308). These results were supported by the later appli- late in anoxic zones due to the activities of sulfate reducers, cation of more sophisticated techniques, in particular flow cy- particularly during periods of low pumping activity (159). Sim- tometry (23, 289, 290). Pile and colleagues reported of ilarly, ammonia and nitrite, which can be toxic to eukaryotes, the Atlantic sponge Mycale lingua on various types of are products of sponge and microbial metabolism but may be (Ͻ10 ␮m in size), with retention efficiencies ranging from 93% oxidized to harmless forms via the activities of nitrifying mi- for Prochlorococcus-type cyanobacteria down to 72% for the croorganisms. While negative effects of nitrite on the develop- smallest photosynthetic eukaryotes (290). Similar methodolo- ment of some juvenile freshwater sponges have been demon- gies applied to the encrusting New Zealand sponge Polymastia

156 Microorganisms in marine sponges

FIG. 19. Effect of a bacterial pathogen on a marine sponge. Transmission electron micrographs of Rhopaloeides odorabile tissue are shown, displaying (A) the diversity of bacterial morphotypes in healthy tissue, (B) a sponge experimentally infected with the alphaproteobacterial pathogen strain NW4327, and (C) consequent necrosis of the sponge tissue. Bar ϭ 500 nm. (Reprinted from reference 455 with permission of the publisher.) croceus showed the best retention of Synechococcus-type cya- lysis of symbionts was in fact an artifact of the histology pro- nobacteria (94%) and (88%), with somewhat cedure (477). Wilkinson argued that while some cyanobacterial poorer retention of Prochlorococcus-type cyanobacteria and cells may be digested intracellularly (e.g., see reference 473), other (noncyano-) bacteria (74 and 46%, respectively) (23). this is the exception rather than the rule. However, several The lower retention of some cell types suggested that P. cro- other reports of bacterial (including cyanobacterial) consump- ceus was selective in its feeding. Laboratory experiments in- tion by sponges have since emerged (172, 222, 266, 422), lend- volving the feeding of symbiotic versus seawater bacteria to ing weight to the notion that certain sponges may “farm” other sponges lend strong support to the notion of selective bacteria as a food source (172). Indeed, phagocytosis and sub- feeding (459, 482). sequent intracellular digestion of bacteria are the presumed The generally highly efficient removal of particles from sea- mechanisms of nutrient transfer between a carnivorous Clado- water is due largely to the extraordinarily large number of rhiza sponge and its methanotrophic symbionts (429, 431). choanocyte chambers (ϳ1 ϫ 107 per cm3) in sponge tissues Harming the host: pathogenesis, parasitism, and fouling. (306). With each chamber containing as many as 150 choano- Deleterious effects of microbes on sponges may be direct (i.e., cytes (371), coupled with the ability of pinacocytes (epithelial pathogenesis or parasitism) or indirect (e.g., microbial films cells) to capture larger particles (308, 412), any food particle promoting surface fouling). The various reported instances of passing through the intricate aquiferous system of a sponge is sponge disease have generally been attributed to bacteria or subjected to intense grazing pressure. Interestingly, it now ap- fungi (199), yet in most cases the responsible microbe(s) has pears that even viruses can be retained by sponges, with some not been identified unequivocally. A notable exception is a 23% of viral particles being removed from seawater by the Red 2002 study by Webster and colleagues (455) in which they Sea sponge Negombata magnifica (131). Considering the enor- isolated a pathogenic alphaproteobacterium (designated strain mous abundance of viruses in seawater (1 million to 100 mil- NW4327) from an infected individual of the Great Barrier lion per ml) (387), this could represent a significant flux of Reef sponge Rhopaloeides odorabile. Strain NW4327, which is nutrients in ecosystems containing large sponge populations. related to the tumor-forming symbionts of Prionitis sp. mac- While most studies of sponge feeding have been conducted roalgae (12) and to the causative agent of juvenile dis- with demosponges (e.g., see references 89, 196, 307, 313, and ease (34, 35), was shown to infect and kill healthy sponge tissue 379), microbial retention efficiencies of 90% or more have also (455). The mechanism by which this occurred was via degra- been reported for hexactinellids (494, 496). The deep-sea dation of the collagenous spongin fibers, with almost the entire hexactinellid Sericolophus hawaiicus was somewhat less effi- sponge surface subject to tissue necrosis following experimen- cient, with microbial retention efficiencies ranging from 47% tal inoculation with strain NW4327 (Fig. 19). Similarly infected for bacteria to 54% for photosynthetic eukaryotes of Ͻ3 ␮m tissue, with documented bacterial attack of spongin fibers, was (291). evident during a devastating outbreak of disease in commer- Consumption of symbiotic microorganisms has also been cially important Mediterranean sponges during the late 1980s raised as a possible food source for sponges (336, 337). The (433). Mass mortalities of commercially important sponges first report of apparent widespread disintegration, both intra- (e.g., Spongia spp.) have occurred several times in both the and extracellularly, of cyanobacterial symbionts was from Sara Mediterranean (199, 298) and Caribbean (119, 199, 375), vir- in the early 1970s (336). His TEM observations suggested that tually eliminating commercial sponge fisheries in some areas. the Mediterranean sponge Ircinia variabilis actively degraded Not only sponges, but also corals and other epibenthic organ- Aphanocapsa-type cyanobacteria (now considered Synechococ- isms, experienced extensive mortality during a 1999 episode in cus spp.) (421) both in the sponge mesohyl and within certain the northwestern Mediterranean (52). This outbreak coincided sponge cells, providing an important source of photosyntheti- with a sudden increase in seawater temperature, with subse- cally fixed carbon to the host sponge (336). These results have quent laboratory studies suggesting the additional involvement since been questioned, with the suggestion that the observed of both protozoans and fungi. Increased microbial virulence

157 Part I

and/or compromised host resistance linked to global warming targets for potentially harmful microorganisms. Compounds has already been postulated as a cause of many mass mortal- with antibacterial, antifungal, or antifouling properties are pro- ities of marine organisms (139, 140), and it will be of great duced by many sponges (19, 32, 112), and those chemicals with interest (and concern) to see how marine sponges are affected more specific effects may allow the host sponge to select for by predicted rises in seawater temperature in the future. Other harmless or even beneficial microbes while deterring deleteri- reports of diseases in sponges include the so-called Aplysina ous types. Interestingly, the resident microbial community may red band syndrome, afflicting aplysinid sponges on Bahaman also participate in host defense, and there are numerous ex- reefs (262), cyanobacterial overgrowth of Geodia papyracea amples of the antimicrobial potential of apparently indigenous (330), and repeated observations of diseased sponges on a microbes (56, 147, 203, 397, 402). In addition, at least one Panamanian coral reef over a 14-year period (492). Bleaching sponge, Halichondria panicea, prevents fouling and sedimen- of Xestospongia muta and other Caribbean sponges has also tary clogging of its ostia by sloughing off its outer tissue layer been reported (64, 91, 251, 441), but it remains to be estab- every few weeks (14), while the innate immune systems of lished whether, as is the case for some corals (325, 326) and an sponges (see “Establishment and Maintenance of Sponge-Mi- Australian macroalga (R. J. Case, A. Low, W. C. Chen, S. crobe Associations”) are also believed to play a role in the Longford, G. R. Crocetti, N. A. Tujula, P. Steinberg, and S. prevention of microbial invasion. Kjelleberg, presented at the 11th International Symposium on Microbial Ecology, Vienna, Austria, 20 to 25 August 2006), The Big Picture: Temporal and Biogeographic Variability in bacterial pathogens are to blame. Bacteria of two genera (Ba- Microbial Communities of Sponges cillus and Pseudomonas) were identified as possible agents of disease in the Papua New Guinean sponge Ianthella basta (54). Variability in sponge-associated microbial communities has This fan-shaped sponge has undergone significant mortality at been examined at a number of levels, such as over time (days a number of inshore sites, leading to speculation that the to months) (105, 201, 390, 462), within and among individuals putative pathogen(s) may be of terrestrial origin. A simple of the same sponge species (mm to thousands of km) (7, 146, model was recently developed to describe the role of sponge 201, 365, 390, 391, 404, 454, 462), and among different host morphology in recovery from disease, with branching sponges species (146, 151, 208, 390). Other studies have investigated being the most likely to recover (493). To our knowledge, the spatial and/or temporal distributions of specific microbial nothing is known about diseases of freshwater sponges. taxa within sponges (100, 226, 294, 421, 453). If an emergent Parasitism of sponges by diatoms has been reported for theme is to be identified from these studies, it is that, with several Antarctic species (16, 51). Bavestrello and coworkers some exceptions, sponge-associated microbial communities ap- found a negative correlation between chlorophyll a (used as a pear to be relatively stable, with little variation in time and proxy for diatom abundance) and sponge carbohydrate levels space (148). The main caveat to this statement is that the (16), while in a parallel study of the hexactinellid sponge Sco- methods employed may not always detect the variability which lymastra joubini, they described a degradation of sponge inter- is present. We review the main published findings on these nal tissue in areas of dense diatom aggregations (51). The topics and, where appropriate, use our own phylogenetic anal- diatoms in S. joubini were of the genus Melosira and appeared yses (see Evolution and Diversity of Sponge-Associated Mi- to enter the host either through the ostia (inhalant openings) croorganisms) to aid our discussion of symbiont biogeography. (Fig. 2) or via active incorporation by the sponge pinacoderm Considering first those studies in which the whole microbial (dermal membrane). Why sponges should actively incorporate community was targeted, only a few papers deal with temporal potentially harmful diatoms is not clear, although consumption variability. The first examined such variation in aquarium- of diatoms as a food source is one possibility (113, 114). Al- maintained Aplysina aerophoba (105). The authors used several ternatively, the silica-encased diatoms may “trick” the sponge methods to characterize the resident microbial community, cells into taking them up (51), a plausible explanation given the including cell counts (both 4Ј,6Ј-diamidino-2-phenylindole tendency of some sponges to incorporate siliceous particles [DAPI]- and cultivation-based), TEM, FISH, and DGGE. from the surrounding environment (15, 17, 18, 107). Currently, What they revealed was an extremely abundant bacterial com- nothing is known about the nature of the interactions between munity (6.4 ϫ 108 cells per g sponge tissue) which varied little sponges and diatoms in tropical and temperate systems (65, during an 11-day incubation period, even under starvation con- 390). ditions or upon exposure to antibiotics. Although DGGE Microbes may also harm sponges in a less direct manner, for banding patterns changed slightly during the antibiotic treat- example, by promoting the fouling of sponge surfaces. Any ment, relative levels of abundance of the major bacterial surface in the marine environment, biotic or abiotic, is subject groups, as assessed by FISH, stayed fairly constant irrespective to intense fouling pressure. During the colonization process, a of treatment (105). The lack of observed temporal variability as new surface will first develop a biochemical conditioning film, well as the apparent resistance of the community to distur- followed by microbial fouling (e.g., colonization by bacteria bance suggests that A. aerophoba harbors a highly stable mi- and diatoms). This biofilm then acts as a precursor to attach- crobial community. Similarly, cultivation for up to 8 months ment by macrofouling organisms, such as invertebrates and did not seem to greatly alter the bacterial community in Geodia macroalgae (71, 450), which in the worst cases can negatively barretti, at least at the broad phylogenetic levels targeted by the affect sponge nutrition by blocking feeding channels or can applied FISH probes (160). Temporal variability has also been increase hydrodynamic drag, resulting in sponge dislodgment examined among sponges in the field. A 16S rRNA gene- from the substratum. DGGE study found bacterial communities in the temperate It is important that sponges not be considered mere helpless Australian sponges Callyspongia sp., Stylinos sp., and Cymbas-

158 Microorganisms in marine sponges

tela concentrica to be highly stable over the course of a year, of large-scale biogeographic variability in sponges was the 2002 while additional sampling of the last species revealed a similar study by Hentschel and colleagues (146), which we discussed at lack of variation on a shorter (days to weeks) time scale (390). length in previous sections. The sponges Rhopaloeides odor- Another DNA fingerprinting method, terminal RFLP analysis, abile (458), Aplysina aerophoba, and Theonella swinhoei con- also identified only relatively minor temporal changes in bac- tained substantially overlapping microbial communities whose terial community composition on the surface of the sponge sequences often fell in monophyletic, sponge-specific clusters, Mycale adhaerens from Hong Kong (201). In contrast to these despite wide (host) phylogenetic and geographic separation studies, the bacterial community profile of the North Sea (146). A 2005 study employed 16S rRNA gene-DGGE to in- sponge Halichondria panicea, as assessed by 16S rRNA gene- vestigate the bacterial community in the sponge Cymbastela DGGE, varied considerably over a 10-month period (462). The concentrica along the eastern Australian coast (391). At eight archaeal community, also assessed by DGGE, varied little. sampling sites spanning 500 km of coastline within the tem- Another study by the same group, on the North Sea sponge perate range of the sponge, bacterial community composition Pachymatisma johnstonia, demonstrated stable bacterial com- varied little. However, C. concentrica sponges from a tropical munities in specimens sampled at different times (2 years location Ͼ1,000 km away had a seemingly very different resi- apart) from two Orkney Isles collection sites (A. Wichels, S. dent bacterial community. Seawater collected during sponge Kuppardt, and G. Gerdts, presented at the 10th International sampling varied comparatively little between the tropical and Symposium on Microbial Ecology, Cancun, Mexico, 2004). temperate locations. Allopatric speciation (resulting from ad- Spatial variability in sponge-associated microbial communi- aptation to geographically separated hosts) is one possible ties has been studied from the millimeter to the interocean explanation for the different communities, although latitudinal scale. Taylor and coworkers examined spatial variability within changes in environmental factors (e.g., temperature and light) and among individuals of three cooccurring Australian sponges could also be responsible. A more prosaic explanation is that (390). In all cases, the variation in 16S rRNA gene-DGGE the C. concentrica individuals from the two locations could banding patterns (and inferred community compositions) was simply be distinct (sub)species, although molecular taxonomy minor, with even the least similar samples for a species sharing studies would be needed to confirm this (391). Cultivation Ͼ70% of bands. For the 30% or less of bands which did vary, efforts by Sfanos and colleagues in which 17,000 bacterial iso- most of the variation could be ascribed to differences among lates were obtained and more than 2,000 were screened by 16S rather than within individual sponges. Considerable differences rRNA-based RFLP fingerprinting and/or sequencing often were seen among different host species, with one sponge in yielded the same bacterium from many sponge hosts from particular harboring a distinct bacterial community compared multiple locations (365). The most extreme case was that of an to those in the other two species (390). In another study, alphaproteobacterium (GenBank accession no. AY362009), Wichels et al. found differences between mesohyl-inhabiting which was recovered from 18 sponge species (plus several microorganisms and transient microbes present in the sponge nonsponge sources, such as a coral and a sea cucumber) spread aquiferous system (462). The latter fraction of microbes was among various Caribbean and eastern Atlantic locations (365). targeted by gently compressing Halichondria panicea tissue Several papers have examined the temporal and spatial vari- within a syringe and collecting the outflowing water. Although ability of particular sponge-associated microorganisms, often incomplete separation of aquiferous system and tissue frac- providing valuable clues to the nature of the sponge-microbe tions may have sometimes disguised differences (462), in gen- association (i.e., true symbionts would be expected to maintain eral there did seem to be distinct communities between the two a long and consistent relationship). For example, “Candidatus sample types. Marked differences were also evident between Cenarchaeum symbiosum,” the sole archaeon present in the outer (cortex) and inner (endosome) tissues in the Mediterra- marine sponge Axinella mexicana, was recorded from all 23 nean sponge Tethya aurantium (404). Cell separation tech- individuals of this sponge collected from the Californian coast niques used in natural product research on sponges have also over a 9-month period (294). Furthermore, archaeal rRNA identified patterns of microbial distribution within sponge tis- levels stayed relatively constant (and high) for more than a sues. For example, cyanobacterial symbionts in the ectosome year in aquarium-maintained A. mexicana, indicating a highly (outer tissues) of Theonella swinhoei were readily separated stable relationship between the sponge and its archaeal inhab- from a filamentous bacterium (later identified as the delta- itant. A subsequent study demonstrated temporally and spa- proteobacterium “Candidatus Entotheonella palauensis” [351]) tially stable associations between three Mediterranean ax- which occurs exclusively in the inner endosome (27, 28). It is inellid sponges (Axinella damicornis, Axinella verrucosa, and typical for phototrophic symbionts, such as cyanobacteria, to Axinella sp.) and their single, host-specific crenarchaeal asso- be prevalent in the outer, well-illuminated surfaces of host ciates (which, in all cases, were related to “Ca. Cenarchaeum sponges, while other microorganisms may dominate the inner symbiosum”) (226). The cultivable fraction of the microbial core (148). community in Rhopaloeides odorabile was always dominated, Moving up to the next spatial scale, we consider geographic irrespective of sampling time or location, by a specific alpha- patterns of variability. A 16S rRNA gene-DGGE study of the proteobacterium (453). This bacterium, which is closely related Antarctic sponges , Kirkpatrickia vari- to Pseudovibrio denitrificans (Fig. 14), comprised Ͼ80% of the alosa, Latrunculia apicalis, Mycale acerata, and Sphaerotylus total heterotrophic bacterial count in samples collected antarcticus revealed that associated bacterial communities over a 460-km portion of the Great Barrier Reef as well as were highly consistent, both among individual sponges at the in those collected during four consecutive seasons at one same sampling site and also among three different sampling reef. Another alphaproteobacterium, affiliated with the genus sites separated by some 10 km (454). The first molecular study Rhodobacter, was found in all samples of the sponge Halichon-

159 Part I

FIG. 20. Global distributions of selected monophyletic, sponge-specific clusters. Symbols refer to collection locations for representatives of the “Ca. Synechococcus spongiarum” (Cyanobacteria) (circles), Actinobacteria (triangles), and Acidobacteria (stars) clusters. In the last cluster, coral-derived sequences from the Mediterranean are also present.

dria panicea from the Adriatic, Baltic, and North (7). In a also widely distributed. The “Poribacteria,” for example, have much earlier study, Wilkinson et al. isolated a particular bac- so far been identified from sponges in the Mediterranean, terium (or at least a highly similar one, as molecular data were Caribbean, and eastern Pacific (100; this study), while a large not feasible at that time) from several sponges in the Mediter- sponge-specific cluster within the Actinobacteria (Fig. 13) con- ranean and the Great Barrier Reef (483). tains sequences from the Red Sea (146), South China Sea The biogeography of sponge-associated cyanobacteria has (data not shown), Indonesia (235), and various Caribbean lo- recently come under close scrutiny. Usher and colleagues per- cations (235, 342; this study). Numerous other such examples formed an extensive survey of cyanobacterial symbionts, sam- exist, indicating an apparently global distribution of many pling nine sponge species (from six genera) in the Mediterra- sponge-specific microbes (Fig. 20). Hill and colleagues (151) nean Sea and the Pacific, Southern, and Indian Oceans (421). attempted to relate the occurrence of major (sponge-associ- In addition, one of these sponges, Chondrilla australiensis, was ated) bacterial taxa to the geographic location where the host sampled from eight Australian locations spanning several sponge was collected. They suggested that some patterns could thousand kilometers and a wide temperature range. 16S rRNA be discerned whereby specific taxa might be better represented gene sequences representing at least four closely related lin- in, e.g., tropical but not cold-temperate sponges. However, we eages of Synechococcus spp. were recovered from the various argue that such statements are premature and that there are host sponges and included, most notably, “Candidatus Syne- insufficient data to draw firm conclusions at present. For ex- chococcus spongiarum” (426), which was present in four of the ample, many (or most) of the existing 16S rRNA gene libraries sampled sponges, including all sampled individuals of C. aus- from sponges have not been sampled exhaustively, and missing traliensis (421). Interestingly, the “Ca. Synechococcus spongia- taxa may well be represented in a library of clones but not yet rum” sequences from the Usher et al. study comprise, together identified due to low sequence coverage. Furthermore, differ- with other sequences obtained independently from the Carib- ent geographic areas (and different habitat types within an bean (78, 151, 342, 383), Red Sea (383), east Africa (383), area) have not been studied equally well, so there could be Micronesia (146, 394), Mediterranean (146), and southeastern biases towards apparently more diversity in some areas which Australia (this study), one of the largest documented mono- have received more attention. The construction of 16S rRNA phyletic, sponge-specific clusters (Fig. 5). In total, “Ca. Syn- gene libraries from sponges in underrepresented locations echococcus spongiarum”-like sequences have been recovered (e.g., Africa, South America, and western ) from 21 sponge species from around the world, making this would go a long way towards improving our understanding of organism similarly widely distributed as its free-living Syne- sponge symbiont biogeography. chococcus/Prochlorococcus counterparts (273, 340). Throughout this review, we have attempted to carefully eval- Many of the sponge-specific clusters from other phyla are uate published data in the context of the methods and ap-

160 Microorganisms in marine sponges

FIG. 21. Chemical structures of jaspamide (left), from Jaspis sp. sponges, and chondramide D (right), from the deltaproteobacterium Chondromyces crocatus. Note the remarkable structural similarities between the compounds.

proaches used for a particular study. This is never more critical BIOTECHNOLOGY OF SPONGE-MICROBE than when considering variability in complex microbial com- ASSOCIATIONS: POTENTIAL AND LIMITATIONS munities. Appropriate sampling designs are important for in- Biologically Active Chemicals from Marine Sponge-Microbe vestigations of any ecological system, yet in the past this has Consortia and Their Commercial-Scale Supply often been overlooked in microbial ecology circles (238, 390). The fact that one should analyze sufficient replicate samples to An enormous number of biologically active compounds have encompass biological variability is beyond question, yet many been isolated from marine sponges and their associated micro- of the available methods for characterizing microbial commu- organisms. Indeed, sponges are the most prolific marine pro- nities are seemingly incompatible with this goal. DNA finger- ducers of novel compounds, with more than 200 new metabo- printing techniques such as DGGE, terminal RFLP analysis, lites reported each year (see reference 32 and preceding and automated ribosomal intergenic spacer analysis offer reviews in that series). Furthermore, more sponge-derived high-throughput analyses of large numbers of samples, but compounds are in clinical and preclinical trials (e.g., as anti- together they suffer from one major drawback: without con- cancer or anti-inflammatory agents) than compounds from siderable efforts (e.g., sequencing of excised DGGE bands), any other marine phylum (31). The occurrence in unrelated the bands or peaks representing particular microorganisms sponges of structurally similar compounds, particularly those have no identity. If banding patterns alone are compared as which were otherwise known exclusively from microorganisms, proxies of community composition, one must acknowledge that led to speculation that such compounds (including some al- observed changes in a community could equally likely be due ready in drug trials) were of microbial origin (27, 143, 280, 427) to changes among two strains of the same microbial species or (Fig. 21). Since chemical synthesis of natural products can be to a much more significant shift, such as from a member of one problematic and expensive due to their structural complexity (4, 48, 373), the realization that at least some compounds may bacterial phylum to another. Conversely, FISH and 16S rRNA be produced by microbes raised hopes of obtaining a sustain- gene library analyses can provide detailed quantitative and able, essentially unlimited supply of compounds for testing and phylogenetic information, respectively, yet neither approach is subsequent drug production (e.g., via cultivation of the rele- well suited to analyzing large numbers of samples. Based on vant bacteria) (280, 297). Today, the true origin of most sponge our own experience as well as reports in the literature (e.g., see compounds has still not been proven unambiguously and re- reference 458), autofluorescence in sponges can also create mains a key issue among marine natural product chemists. The difficulties for FISH analyses. A need therefore exists for a possibility of of biosynthetic pathways phylogenetically informative yet rapid means of assessing mi- among different sponges has also been raised (332). It is not crobial community structure. Microarrays offer particular our intention to comprehensively review sponge-derived natu- promise in this regard, with a range of 16S rRNA- and func- ral products; such reviews are the subject of chemistry rather tional gene-based microarrays already available (reviewed in than microbiology per se, and many excellent reviews dedi- references 122 and 215). The highly parallel nature of microar- cated to this topic already exist (31, 32, 191, 236, 279, 280). rays provides the potential, for example, to survey the presence Rather, we focus our attention on selected important examples of multiple sponge-specific clusters in a single assay, something and highlight some of the difficulties involved with obtaining a which is not possible with other existing techniques. Impor- consumer-ready end product. tantly, symbiont function could also be addressed via the Sponge (or microbe)-derived compounds span a wide range so-called isotope array approach (2). of chemical classes (e.g., terpenoids, alkaloids, , and

161 Part I

FIG. 22. Chemical structure of halichondrin B. polyketides) with an equally wide range of biotechnologically chondrins has never been determined unambiguously). Lisso- relevant properties (e.g., anticancer, antibacterial, antifungal, dendoryx sp., collected from the coast of southern New Zeal- antiviral, anti-inflammatory, and antifouling) (31, 32, 112, 186, and, yielded the largest amounts of halichondrins and 229, 236, 279, 280). The attention of natural product chemists therefore became a focus of drug supply efforts (138, 250). and pharmaceutical companies, at present, is focused firmly on Based on the potency of halichondrin B and its projected anticancer drugs, with several promising sponge-derived com- demand if approved for human use, the requirement for clin- pounds in clinical and preclinical cancer trials (31, 255, 370). ical trials was estimated to be ϳ10 g, with annual requirements The large number of novel, active metabolites being reported as a commercial drug of 1 to 5 kg (138). Given that 1 metric ton from sponges every year begs the question of why such chem- of Lissodendoryx sp. sponges yielded only 300 mg of halichon- icals have not yet made it to pharmacy shelves. To date, and to drin B and that the entire natural biomass of the sponge was the best of our knowledge, not a single compound obtained estimated to be only 289 metric tons, collection from the wild from a sponge has been approved as a drug, with a major brake was quickly ruled out. Aquaculture of Lissodendoryx sp. was on progress being the so-called supply problem (138, 297, 408). then investigated, with promising initial results (250). How- (The nucleoside analogs Ara-A and Ara-C, commercialized as ever, scale-up of the operation to the levels necessary for antiviral and anticancer agents, respectively, could arguably be commercial production was not achieved due to a lack of considered the sole exceptions. They were not isolated directly funding (M. J. Page, personal communication), and the small from sponges but are synthetic derivatives based on com- amounts of compound present in the sponge tissue may render pounds from the Caribbean sponge Cryptotethia crypta [24, the aquaculture option economically untenable in this case 25].) Biologically active natural products are often produced in anyway (373). Nevertheless, halichondrin B may yet prove to relatively small amounts, and often by rare animals whose be a success story, with a synthetic analog, E7389, in phase I natural populations cannot sustain the extensive collections clinical trials as an anticancer compound (370). This simplified required for clinical trials. Alternative means for producing version of halichondrin B is more amenable to chemical syn- large amounts of metabolites are therefore required. We illus- thesis but retains the biological activities of the original com- trate this issue by using two examples, the anticancer com- pound (60). pounds halichondrin B and peloruside A. Our second example concerns the macrocyclic lactone The halichondrins are a group of polyether macrolides that peloruside A (460) (Fig. 23). Isolated from the New Zealand exhibit potent antitumor activities (158, 415). First isolated demosponge Mycale hentscheli, peloruside A shows promising from the Japanese sponge Halichondria okadai in the mid- 1980s (158), they were subsequently found in several other sponges from diverse geographic locations, including Axinella spp., Phakiella carteri, Raspailia agminata, and Lissodendoryx sp. (138). Halichondrin B (Fig. 22) was particularly sought after due to its high cytotoxicity, and its total synthesis was reported as early as 1992 (4). However, due to the structural complexity of the compound, many steps were required for its synthesis, rendering total synthesis impractical for industrial- scale production. While the occurrence of halichondrins in many unrelated sponges suggested a microbial origin, little was known about the microbiology of the relevant sponges, and thus alternative avenues were investigated (to our knowledge, the precise [i.e., sponge versus microbial] origin of the hali- FIG. 23. Chemical structure of peloruside A.

162 Microorganisms in marine sponges

anticancer properties, acting in a similar manner and potency Cultivation of metabolite-producing microorganisms. Culti- to the widely used cancer drug paclitaxel (Taxol) (166). With vation of sponge-associated microorganisms that produce bio- the compound currently in preclinical trials, two avenues are active compounds is the most direct method for large-scale being pursued in parallel to ensure a sufficient supply of the production of these chemicals (154), and cultivation ap- compound for subsequent clinical trials. Chemical synthesis is proaches are widely practiced among those targeting bioactive one approach, with several groups recently reporting partial or compounds (46, 47, 81, 130, 147, 154, 163, 176, 189, 235, 364, total synthesis (e.g., see reference 177). A New Zealand con- 365, 378, 453). The potential payoffs from the cultivation ap- sortium, working together with a U.S. pharmaceutical com- proach are obvious and substantial: if metabolite producers pany, is currently investigating whether cost-effective, industri- can be isolated on artificial media and grown to significant cell al-scale synthesis is achievable (137). An alternative supply numbers (while continuing to produce the relevant metabo- option for peloruside A is being explored by the same group, lite), then this obviates the need for large-scale harvesting of with aquaculture of M. hentscheli looking highly encouraging natural sponge populations, with its environmentally and finan- (137, 271). With 200 kg of sponge yielding a mere2gofpure cially negative implications. peloruside A, scaling-up is a priority, with the goal of growing Two broad strategies for isolating microbial producers of Ͼ500 kg of sponge over the coming year (137). Other com- bioactive compounds were outlined by Hill in a recent review pounds of pharmaceutical interest are also produced by M. (154). The first is to use a wide range of media in an effort to hentscheli, namely, the cytotoxic polyketide mycalamide A and grow as many different sponge-associated microbes as possible. the macrolide pateamine (165, 256, 270, 278). Concentrations Since growth under different culture conditions may influence of these metabolites in natural sponge populations vary signif- which metabolites are produced, the use of many different icantly in time and/or space (270), suggesting that complex media and conditions should help to maximize the chemical ecological and physical factors may be involved in their pro- diversity from a given microorganism (154). Bacteria associ- duction. An improved understanding of the ecological roles of ated with deep-sea benthic invertebrates have been the subject these and other compounds could greatly benefit metabolite of extensive cultivation efforts by the Harbor Branch Oceano- harvesting programs, and indeed, ecological observations are graphic Institution, with a range of nutrient-poor to nutrient- often used to guide the initial stages of drug discovery in rich media being utilized (130, 264, 365). Approximately 17,000 marine environments (295, 359). Ongoing microbiological in- isolated microbes, most from deep water and mostly from vestigations with M. hentscheli (452; S. A. Anderson, unpublished sponges, are present in the Harbor Branch Oceanographic data) should also benefit future drug development efforts with Marine Microbial Culture Collection (365). These include rep- this sponge. resentatives of the Proteobacteria, Bacteroidetes, Firmicutes, Supply issues notwithstanding, the pharmacological po- and Actinobacteria and are the subject of natural product tential of marine sponges and other sessile invertebrates screening. An alternative, more targeted approach is to go (e.g., corals, bryozoans, and ascidians) is enormous. Although after specific microbial groups with proven track records in the progress toward the commercial product stage has been slow, production of bioactive compounds. Many such groups, includ- it is highly likely that at least one of the several compounds ing cyanobacteria, fungi, and actinomycetes, are well known now in clinical trials (or a synthetic analog) will be commer- from sponges (148, 163), with actinomycetes being the subject cialized within the next few years. A combination of improved of a particularly interesting success story. Sponge-derived chemical synthesis methods with the various approaches out- actinomycetes of the genus Micromonospora produce man- lined in the following section should ensure a bright future for zamines, alkaloids with, among other things, potent antimalar- this field, with sponge-derived natural products being utilized ial properties (10, 93, 155, 301). The first hint that manzamines either in their natural form or as inspiration for new, labora- were of microbial origin came from the finding of these com- tory-generated compounds (e.g., via chemical proteomics) pounds in many distantly related, geographically disparate (287). As a footnote to this discussion, the freshwater sponges sponge species. Subsequent cultivation-dependent and -inde- should also be mentioned. Their chemistry has received much pendent characterization of the microbial communities in less attention than that of their marine counterparts, and while two Indonesian manzamine-producing sponges, 01IND 35 and various and a compound with antipredator activity have 01IND 52, revealed highly diverse assemblages, with the re- been reported (77, 311), it is unclear whether these sponges covery of actinomycetes provoking intensive culturing efforts produce many, if any, compounds of pharmaceutical interest. in their direction (154). Growth of the sponge-derived Mi- cromonospora sp. has since been achieved on a large scale in Methods for Accessing the Hidden Chemistry 20-liter , with maintenance of manzamine pro- of Marine Sponges duction (R. T. Hill, personal communication). Improvements in this process should greatly facilitate passage of these com- A number of (nonsynthesis) approaches are available for pounds through the various stages of the drug-testing process. accessing biologically active natural products from sponges and Actinomycete-selective media were also used successfully with the microorganisms within them (Fig. 24). For convenience, we the sponges Pseudoceratina clavata (188), Xestospongia spp. split these into the following three main themes: cultivation of (235), Hymeniacidon perlevis (498), and Craniella australiensis metabolite-producing microbes, sponge culture, and molecular (209). In the last study, many of the cultivated actinomycetes biological methods, such as metagenomics. In addition, we displayed broad-spectrum antimicrobial activities. highlight the importance of metabolite localization studies for There are numerous other examples of the production of improving our knowledge of which partner (sponge or micro- biologically active compounds by sponge-derived microbial iso- organism) is responsible for metabolite production. lates. An antibacterial was isolated from both the

163 Part I

FIG. 24. Approaches for obtaining bioactive metabolites from marine sponges.

sponge Hyatella sp. and an associated Vibrio sp. (260), while a reisolation from the original sponge hosts. Interestingly, it ap- glycoglycerolipid produced by a Halichondria panicea-derived pears that unlike bacteria, fungi are not the source of any Microbacterium sp. had antitumor properties (463). In another natural products previously ascribed to marine sponges (191). study, several quinolones, including one with both antimicro- The success of efforts to isolate sponge-associated microor- bial and cytotoxic activities, were isolated from a pseudomonad ganisms that produce bioactive compounds is dependent upon from the Pacific sponge Homophymia sp. (45). Although the a number of factors. Most significantly, the majority of envi- mechanistic basis was not identified, 27 bacteria isolated from ronmental microorganisms, including those in sponges, have the Mediterranean sponges Aplysina aerophoba and A. caver- proven resistant to cultivation by standard techniques (105, nicola exhibited antimicrobial activities in a series of assays 380). Although various authors have reported improved cul- (147). Given the activity of some of these isolates against turability of sponge-associated bacteria via, for example, sup- clinically important multiresistant Staphylococcus aureus and plementing the media with sponge tissue extracts (458) or Staphylococcus epidermidis strains, assay-guided fractionation and sodium pyruvate (264), the proportion of total and subsequent chemical characterization of any active com- sponge bacteria that can be isolated has remained low. Only ponents could prove particularly profitable. Terrestrial fungi 0.06, 0.1, 0.15, and 0.7% of total bacteria could be cultured have a long-standing reputation as prolific producers of bioac- from the sponges Candidaspongia flabellata (47), Rhopaloeides tive natural products (44), and it is hardly surprising that odorabile (47, 453), Aplysina aerophoba (105), and 01IND 35/ sponge-associated fungi also show promise in this regard. 01IND 52 (154), respectively. Santavy and colleagues were able Many examples exist of sponge-derived fungi that produce to achieve somewhat better recovery (3 to 11%) from the bioactive compounds (29, 42, 44, 163, 174, 176, 295, 296, 451). Caribbean sclerosponge Ceratoporella nicholsoni, although ob- Although many of the isolated fungi are of suspected terres- viously, even in this case, some 90% of the resident bacteria trial origin (i.e., they are closely related to typical terrestrial were not captured by cultivation attempts (335). While culti- species), to some extent this does not matter for drug discovery vation difficulties are hardly confined to sponges, it is never- purposes; even if sponges act only as mere accumulators of theless likely that many of the sponge associates are obligate contaminant fungi, these microorganisms can still be targeted, symbionts which may have evolved with the sponge over hun- and once they are isolated, there may be no need to attempt dreds of millions of years (see Evolution and Diversity of

164 Microorganisms in marine sponges

stepped by cultivating sponges under semienclosed or even fully closed conditions (22, 90, 141, 161, 267–269). Although this is generally more expensive than sea-based aquaculture, an obvious advantage is the ability to control environmental pa- rameters, such as growth temperature, water movement, and food supply, as well as to eliminate biomass loss due to storms or disease outbreaks (see “The Varied Nature of Sponge- Microbe Interactions”). Potential problems in recirculating systems include a buildup of toxic secondary metabolites and metabolic wastes, such as ammonia (22). Considerable growth (Ͼ200% in 1 to 2 months) of Pseudosuberites andrewsi explants was achieved in a bioreactor (268), with even more growth (Ͼ1,000% in 45 days) observed for the sponge Crambe crambe in a closed system (21). Like the case for sea-grown sponges, metabolite production has been observed for sponges sub- FIG. 25. In-sea aquaculture of the Great Barrier Reef sponge Rho- jected to in vitro cultivation (e.g., see references 75 and 90). paloeides odorabile. (Image courtesy of Rocky de Nys [James Cook However, one must remember that despite its promise, this University, Australia], reproduced with permission.) technology remains in its infancy, and to our knowledge, there are no examples of industrial-scale in vitro cultivation of sponges. Sponge-Associated Microorganisms) and will, due to nutri- Sponge cell culture for the production of biologically active tional or other dependencies, be extremely difficult to obtain in metabolites represents the other extreme of the scale contin- pure culture. Furthermore, those that can be isolated may not uum from sea-based aquaculture. Although sponge cells can be necessarily produce the compound anymore, as they may re- dissociated readily and even induced to divide in suspension quire some as yet unknown cue or metabolic intermediates for several cycles, it has so far proven impossible to establish from the host sponge. Additionally, and for unknown reasons, continuous cell lines (reviewed in references 293, 319, and some bacteria simply stop producing the compound of interest 320). Primmorphs, which are three-dimensional aggregates after a certain time on artificial media (145). This could be due comprising proliferating and differentiating sponge cells, have to any of a number of genetic reasons relating to a lack of therefore generated much recent interest since they can be selective pressure in pure culture, e.g., point mutation in a key maintained for long periods (66, 240, 242, 247, 293). Particu- gene or loss of a mobile genetic element carrying the biosyn- larly exciting was the finding by Mu¨ller and coworkers that thesis genes. primmorphs from Dysidea avara grown in a bioreactor pro- Sponge culture. Culturing sponges is another way to address duced the secondary metabolite avarol, which is both charac- the supply problem, irrespective of whether compound pro- teristic of this sponge and of great pharmacological interest duction is due to the sponge or the symbiont. Methods em- due to its strong biological activities (e.g., antitumor, antibac- ployed for the cultivation of sponges for drug production vary terial, and antiviral activities) (242). In contrast, single D. avara widely in scale and sophistication, from sea-based aquaculture cells did not produce avarol. Another interesting aspect of to in vitro cultivation (in closed or semiclosed systems) to cell primmorphs, especially within the context of this review, is that and tissue cultures (22). The technical and economic potential symbiotic microorganisms can be retained within them (247, of each of these was reviewed recently (373). 398), potentially allowing for primmorph production of both In-sea sponge aquaculture has received considerable atten- sponge- and microbe-derived compounds. Since their initial tion for its potential to cost-effectively address the metabolite demonstration in Suberites domuncula (66, 247) and then D. supply issue (Fig. 25) (84–88, 132, 137, 231, 248, 271, 299, 439). avara, primmorphs have been generated from a wide range of Of all approaches, it has the advantage of most closely simu- sponges, including Axinella polypoides, Cliona celata, Halichon- lating the conditions encountered by sponges in nature, and dria panicea, Petrosia ficiformis, and Stylotella agminata (374, practitioners have been able to draw upon more than a century 434, 500). In the coming years, it should become clear whether of experience in farming bath sponges. On the negative side, primmorphs can be scaled up sufficiently to overcome the the inherent unpredictability of marine environments can cre- supply problem for many promising drug leads. ate problems (e.g., due to atypical climatic conditions or storm Surprisingly little information exists on the microbiology of damage) (439) which are easily avoided in controlled in vitro cultured sponges (in any system), yet this could be of vital systems. The outcomes of sponge aquaculture trials have var- importance if metabolites are produced by microbial associ- ied widely, with success dependent upon a number of factors, ates. For example, if sponges are cultured away from their including the type of farming structure (84, 85, 132), sponge natural environment, then metabolite-producing symbionts growth form (86), farming location (271), and season of trans- may conceivably be lost, or if metabolites are diet derived (as plantation (86, 87). If sponge survival can be ensured, then suspected for okadaic acid) (297), then a change in diet would growth increases of up to 5000% per year (relative to the presumably result in a loss of compounds. Moreover, even if starting size) are achievable, depending on the sponge species the desired metabolite is produced by the sponge itself, micro- examined (271). Crucially, bioactive metabolites are typically bial symbionts may still be of direct (e.g., by providing meta- retained in farmed sponges (84, 87, 132, 248, 250, 271). bolic precursors) or indirect (e.g., by affecting general sponge The consequences of environmental variability can be side- health) significance (154).

165 Part I

Metagenomics. One of the most exciting developments in bases of pederin biosynthesis, Piel et al. investigated the pro- molecular biology from a drug discovery perspective has been duction of onnamide in the sponge Theonella swinhoei from the advent of environmental genomics, or metagenomics (92, Japan (285). Ketosynthase (KS) fragments were PCR ampli- 104, 136, 207, 346, 438). Metagenomics refers to the analysis of fied directly from the sponge metagenome, revealing a di- genome fragments from a complex microbial community and verse range of sequence types. More importantly, PCR-based offers the potential for large-scale, sustainable production of screening of a 60,000-clone cosmid library with the same prim- bioactive metabolites, including those produced by unculti- ers yielded a single KS-positive clone, which was fully se- vated microorganisms. If biosynthesis genes can successfully be quenced. Strong indications existed for a bacterial origin of this cloned and expressed in another (cultivated) microorganism, genome fragment (e.g., a lack of introns and small intergenic such as E. coli, then this could ensure an unlimited supply of a distances), which should correspond to almost the entire re- specific metabolite (48, 143). Different approaches for the gion of the polyketide structure needed to obtain an antitumor cloning and heterologous expression of biosynthesis genes compound (285). In addition to the obvious biotechnological from marine invertebrate symbionts were recently the subject importance of this study, as heterologous expression of such of a comprehensive review by Hildebrand and colleagues gene clusters could lead to an inexhaustible supply of target (149). The successful application of many of these methods is metabolites for clinical trials, interesting ecological and evolu- exemplified by studies by Haygood and coworkers on the sym- tionary questions were also raised. For example, how did such biosis between the bryozoan Bugula neritina and its bryostatin- similar biosynthetic pathways come to be present in symbionts producing gammaproteobacterial symbiont, “Candidatus En- of such dissimilar hosts, i.e., marine sponges and terrestrial dobugula seritina” (69, 70, 142, 143, 149, 150). Here we focus beetles? our attention on the results of recent metagenomic studies PKSs have also been studied in other sponges by using meta- involving sponges. genomic approaches. In an attempt to isolate genes encoding Two main types of analysis have been used to extract bio- the promising antitumor compound discodermolide from the technologically relevant information from metagenome librar- Caribbean sponge Discodermia dissoluta, Schirmer and col- ies: one is based on function, whereby libraries are screened for leagues (342) employed a two-step approach. First, degenerate the expression of specific traits, and the other is based on KS primers were used to amplify 256 sequences (85 different screening for sequences themselves (346). While screening for KS sequences), including several from trans-AT-type PKS do- functional traits (e.g., antibiotic production or quorum-sensing mains of the pederin and onnamide types. A selection of the inhibitors) has been successful (to various degrees) in other derived sequences was then used to create a probe pool for the environments (68, 318, 489), studies of sponge metagenomics, screening of 155,000 macroarrayed fosmid and cosmid clones; to our knowledge, have been exclusively sequence based. To given the proportion of bacterial inserts in the studied libraries date, the major foci of such studies have been the polyketide (Ͼ90%) and the average insert size (35 kb), Ͼ4 Gb (some synthase (PKS) and nonribosomal peptide synthetase genes 1,000 bacterial genome equivalents) were screened (342). In (187, 284, 285, 342). The PKSs are responsible for the synthesis total, 1,025 PKS-positive clones (0.7% of all analyzed clones) of bacterial polyketides, a diverse group of pharmacologically were identified. Interestingly, sequencing of selected fosmid important natural products which include the antibiotics eryth- and cosmid clones revealed suprisingly little overlap between romycin and tetracycline as well as antitumor, immunosuppres- these KS domains and those derived from the same sample by sive, and cholesterol-lowering agents (197). Type I PKSs are the direct PCR approach. A PKS consistent with the biosyn- organized in a modular fashion, lending themselves to the thesis of discodermolide was also not found (342). Direct am- combinatorial biosynthesis of novel polyketides with poten- plification of KS domains was also combined with the construc- tially useful pharmaceutical properties (197). A number of tion of fosmid libraries to study PKSs in another sponge, the marine drug candidates, including the bryostatins, disco- Great Barrier Reef species Pseudoceratina clavata (187, 188). dermolide, and the aforementioned peloruside A, belong to Each approach led to the retrieval of five KS domains, all of this class of compounds (104, 150). The modular nature of the which fell into an apparently sponge-specific KS cluster (to- polyketides suggests that environmentally retrieved PKS frag- gether with sequences obtained from Discodermia dissoluta, ments, which may not produce intact bioactive compounds, Theonella swinhoei, and an unidentified sponge) following phy- could still be useful by providing modules for combinatorial logenetic analysis (187). Cultivated bacteria from P. clavata polyketide synthesis (187, 197). A large part of the sponge-PKS were also screened by PCR for KS domains, with 10 such story comes from work by Piel and colleagues (279–286). Their domains detected in representatives of the Actinobacteria, study of the polyketide onnamide in sponges was greatly facil- Alphaproteobacteria, and Firmicutes. None of the KS do- itated by prior metagenomic investigations into the production mains from isolates clustered with the metagenome-derived of a structurally highly similar antitumor compound, pederin, sequences, highlighting the importance of a polyphasic ap- in beetles of the genus Paederus (281, 283, 286). The microbial proach to encompass as much of the PKS diversity as possible community within these beetles is much less complex than that (187). of sponges, allowing easier access to the genome of the bacte- Although not part of a drug isolation strategy per se, no rial pederin producer (349). Ultimately, pederin production discussion of sponge metagenomics would be complete without was linked to a beetle symbiont closely related to Pseudomonas considering the seminal work of DeLong and colleagues (134, aeruginosa, although evidence for lateral gene transfer of the 135, 294, 343–345). Over the past decade, the symbiosis be- pederin-type genes suggests that compound production may tween the Californian sponge Axinella mexicana and the psy- not necessarily correlate with rRNA-based bacterial phylogeny chrophilic crenarchaeote “Candidatus Cenarchaeum symbio- (283, 285). Armed with a sound understanding of the genetic sum” (294) has been a model system for environmental

166 Microorganisms in marine sponges

genomics. “Ca. Cenarchaeum symbiosum” was the first known proaches to revolutionize natural product research with symbiotic crenarchaeote, and despite its as yet uncultivated sponges is undeniable, yet there remain considerable technical status, several factors have made it a suitable target for challenges. For example, if the microbial communities under genomic studies: it is the only archaeon in the sponge and study are highly complex, then the genomes of target organ- dominates the microbiota, consistently comprising some 65% isms (e.g., “Poribacteria”) may remain largely hidden against a of all prokaryotic cells; it is always associated with the sponge; background of genomes from other symbionts (but see Con- and the symbiosis is maintained for a long time in aquaria clusions and Future Directions for possible means of enriching (294). Relatively large amounts of biomass and DNA have specific genomes). Other potential problems include the use of therefore been available, with physical enrichment for the ar- inappropriate host organisms for expression studies (some chaeal cells greatly facilitating the construction of large-insert hosts may not express the genes of interest) (104, 136) and the genomic libraries for this organism. The first published results large sizes of many gene clusters, which can prohibit their from the genomic analyses outlined the characterization of a successful cloning (although there exists the theoretical possi- DNA polymerase which was homologous to those of cultivated bility of reconstructing biosynthetic pathways via the assembly archaeal hyperthermophiles and yet, as revealed by heterolo- of many smaller, overlapping sequence reads). In still other gous expression in E. coli, was inactivated at temperatures cases, nonclustering of biosynthesis genes may be a problem, above 40°C, reflecting the symbiont’s low-temperature lifestyle with the genes of interest spread across different parts of the (345). Initial studies based on the 16S rRNA gene indicated symbiont’s genome (281). On the bright side, the relevant the presence of a single archaeal phylotype in A. mexicana technology is developing quickly. Massive sequencing efforts, (294), so subsequent genome-derived indications of the pres- such as the recent Sargasso Sea study (440), may provide one ence of two closely related variants were unexpected (343). means of accessing rare genomes and the biotechnologically Although the two variants differed Ͻ0.8% in their 16S and 23S relevant information within them. Indeed, an ambitious small- rRNA genes and had an identical gene order for a studied and large-insert sequencing study along these lines was re- 28-kb region, variations in DNA identity of up to 20% were cently initiated for a temperate Australian sponge (S. Kjelle- observed for protein coding regions, with up to 30% variation berg, P. Steinberg, and T. Thomas, personal communication). for intergenic regions (343). These findings thus highlighted Similarly, the new pyrosequencing technology from 454 Life the difficulties created by genomic microheterogeneity in as- Sciences (227) has already been applied successfully to com- sembling environmentally retrieved genome fragments, and plex environmental samples (204) and may prove valuable in these difficulties remain today (76, 414, 440). Despite such future sponge metagenomics projects. Furthermore, high- obstacles, the DeLong group was able to assemble a closed throughput screening techniques have developed to the extent genome for “Ca. Cenarchaeum symbiosum” (134), which has that a single worker can now screen, from start to finish, a already yielded important insights into the metabolism of the library of 400,000 clones for a particular gene in a mere 4 days sponge symbiont in particular and of marine Crenarchaeota in (169). general (135). For example, genome reconstruction revealed Cell separation and metabolite localization. The aforemen- the potential of “Ca. Cenarchaeum symbiosum” to function tioned techniques each offer the potential to generate large either autotrophically (as an ammonia oxidizer) or mixotrophi- quantities of biologically active metabolites from sponge-mi- cally. In an extension of the environmental genomics approach, crobe associations. All three approaches (microbial cultivation, homologues of genes involved in carbon and nitrogen metab- sponge culture, and metagenomics) can, in principle, be un- olism were also found in metagenome libraries from ocean dertaken without prior knowledge of which partner (sponge or waters worldwide, demonstrating the ubiquity of these meta- microorganism) produces a given compound. However, the bolic pathways among marine crenarchaeotes (135). Con- most rational approach is to first establish which cell type(s) is versely, certain genes encoding cell surface, regulatory, or de- responsible for metabolite production, as this can determine a fense mechanisms were not recovered from the free-living logical strategy for future efforts (Fig. 24). Many researchers relatives of “Ca. Cenarchaerum symbiosum,” suggesting that have employed cell separation techniques for this purpose (27, these could be involved specifically with the establishment and 28, 99, 103, 120, 314, 315, 332, 416, 417). A major breakthrough maintenance of the symbiosis (134). in this area came with the realization that chemically fixed The organism-oriented approach taken for “Ca. Cenar- sponge and microbial cells retain their natural products in a chaeum symbiosum” was also employed in a recent study of form amenable to chemical analyses, such as high-performance the sponge-specific candidate phylum “Poribacteria” (100, liquid chromatography and nuclear magnetic resonance spec- 101). Virtually nothing is known about the physiology and troscopy (416). Coupled with the relative ease with which one genetic makeup of these bacteria, and since there are no can dissociate sponge tissue, this opened up a number of pos- cultured representatives, metagenomics offered a promising sibilities for natural product research. In the 1990s, researchers approach. The sole entry point into the “Poribacteria” ge- in Faulkner’s laboratory thus took advantage of cyanobacterial nome was the 16S rRNA gene sequence, and a single fosmid autofluorescence to separate, via flow cytometry, the filamen- clone among 29,000 clones (corresponding to 1.1 Gb of tous cyanobacterium Oscillatoria spongeliae from sponge and DNA in total) was positive in an initial 16S rRNA PCR- other microbial cells in dissociated tissue of Dysidea herbacea based screening (101). Analysis of this 39-kb insert revealed (416, 417). Of three major types of metabolites known previ- 27 open reading frames, including one encoding a new kind ously from this sponge, two—polychlorinated peptides (416) of molybdenum-containing oxidoreductase and several en- and polybrominated biphenyl ethers (417)—were found exclu- coding unusual transmembrane proteins (101). sively inside O. spongeliae cells. In contrast, sesquiterpenoids The potential of metagenomics and other cloning ap- were confined to the sponge cells. While autofluorescing cells

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(e.g., cyanobacteria) have been separated from other cell types production in that cell or, indeed, even in that organism at all. (e.g., sponge and noncyanobacterial microbes) by flow cytom- A compound may diffuse or be exported away from its site of etry, other characteristics, such as cell size and, provided the synthesis, especially if it is toxic to the producer (149). The natural products are retained, FISH probe-conferred fluores- recent demonstration of swinholide A production by free-living cence, could also be used to differentiate sponge and symbiont marine cyanobacteria (9) is a case in point: while Bewley and cells by this technique. coworkers (28) found this compound only in heterotrophic A more common approach has been to separate different bacteria (outlined above), it now seems plausible that cya- cell types in dissociated sponge tissue via centrifugation (27, nobacterial symbionts in Theonella swinhoei may produce the 28, 314). Again, the Faulkner group was instrumental in estab- compound but excrete it to be stored elsewhere. Alternatively, lishing these approaches for sponge natural product study. In both the heterotrophic bacteria and the cyanobacteria may the chemically rich lithistid sponge Theonella swinhoei, Bewley share the capacity for swinholide A biosynthesis, perhaps due and colleagues (28) identified four distinct cell types by elec- to lateral gene transfer (9). Interphylum and even interdomain tron microscopy (sponge cells, unicellular cyanobacteria, uni- transfer of natural product pathways has been reported a num- cellular heterotrophic bacteria, and filamentous heterotrophic ber of times (e.g., the ␤-lactam biosynthesis genes, responsible bacteria [the term “heterotrophic bacteria” is used here in for the production of antibiotics, including penicillins and reference to noncyanobacterial microorganisms; these cells cephalosporins, are found in both fungi and bacteria [41]). Yet could, in principle, be from chemolithoautotrophic bacteria]). another potential explanation is that a given metabolite could By following gross dissection of the sponge (into the endosome be derived from the sponge’s diet. There is evidence suggesting [inner tissues] and ectosome [outer tissues]) with dissociation that this may be the case for okadaic acid, which has been and centrifugation, they were able to obtain relatively clean isolated from various sponges but is known to be produced by fractions comprising each cell type. The macrocyclic polyketide free-living marine dinoflagellates (297, 467). Interestingly, cytotoxin swinholide A was found only in the fraction contain- while localization studies have very often implicated the host ing the unicellular heterotrophic bacteria, while the potent sponge as the source of bioactive metabolites, in at least some antifungal glycopeptide theopalauamide (350) was associated cases it appears likely that intracellular or cell surface-associ- with a filamentous bacterium (later identified as a deltapro- ated bacteria may have been present but overlooked (149). In teobacterium, “Candidatus Entotheonella palauensis” [351]) such cases, microbial production of metabolites cannot be (28). Density gradient centrifugation provides yet another op- ruled out completely. portunity for separating sponge and microbial cells. In this It is important to note that while any of the aforementioned case, dissociated sponge tissue is placed above a density gra- localization techniques may give an indication of which is the dient (consisting of, e.g., Ficoll or Percoll), and following cen- metabolite-producing organism, none of them directly allows trifugation, various cell fractions are formed based on differing the harvesting of large amounts of the desired compounds (in densities (103, 120, 121, 315). Individual fractions can then be contrast to cultivation of the relevant microorganism, sponge examined chemically and microscopically to correlate the oc- culture, or heterologous expression of biosynthesis genes). The currence of compounds with specific cell types. For Haliclona chief benefit of such studies is therefore to improve our un- sp., Garson and coworkers used Percoll gradients to demon- derstanding of metabolite production within the sponge-mi- strate localization of the cytotoxic alkaloids haliclonacyclamine crobe association, identifying the putative producer(s) and thus A and B within the sponge cells rather than in associated providing the basis for targeted cultivation and/or molecular dinoflagellates (120). approaches. Depending on the properties of and prior knowledge about the compounds under study, localization may be achieved with- Other Biotechnologically Relevant Aspects of Sponges out the need for prior cell separation. For example, bromi- nated compounds, which are often biologically active and can In addition to the by now well-known pharmacological po- be present in large amounts in sponges, may be visualized in tential of marine sponges and their associated microbiota, sev- situ by X-ray energy-dispersive microanalysis (406, 410). Spe- eral other aspects are also worth highlighting. For example, the cific antibodies for the toxin latrunculin B were used to prove farming of certain sponges, particularly those in the genera its localization in sponge but not symbiont cells within the Red Spongia and Hippospongia, for their use as bath sponges has Sea demosponge Negombata magnifica (125). In a novel ap- been going on for more than a century. The industry, based proach for sponges, catalyzed reporter deposition-FISH was mainly in the Caribbean and Mediterranean, has been hit recently used to demonstrate mRNA expression from the numerous times by mass outbreaks of sponge disease (see dysB1 genes (responsible for the biosynthesis of polychlori- “Harming the host: pathogenesis, parasitism, and fouling”). A nated peptides) in cells of the cyanobacterium Oscillatoria better understanding of sponge microbiology in general, and spongeliae in Dysidea (Lamellodysidea) herbacea (102). The disease ecology in particular, should contribute to the future biological result was consistent with earlier work by others success of these endeavors. The enormous filtering capacity of (103, 416), but more significantly, this study proved the utility sponges has led to the suggestion that they be farmed in a of the method for the investigation of target chemicals in bioremediation context, e.g., to reduce the high bacterial loads sponges. The main limitation of the method is that a certain resulting from sewage discharges and marine fish farms (110, level of genetic information must first be available for the 233, 379). Such an approach could be optimized financially by studied, or a related, biosynthetic pathway. farming sponges that, in addition to their role as a biofilter, are Although strongly suggestive, even the localization of a me- useful as bath sponges or produce bioactive metabolites which tabolite to a particular cell type is not unequivocal proof of its can subsequently be harvested (233).

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Another aspect relates to the remarkable structural proper- rent paucity of knowledge about many of the microbial asso- ties of the silica of demosponges and hexactinellid ciates of sponges. For many, or even most, symbionts in (glass) sponges. A series of recent studies examined in detail sponges, all that is known so far is their 16S rRNA gene the siliceous spicules of Euplectella aspergillum, a deep-sea sequence: their metabolism remains a black box. Despite var- hexactinellid from the western Pacific (5, 6, 385). Individual ious caveats, the 16S rRNA data assembled here, together with spicules exhibit, in addition to their role in providing structural analyses of so-called functional genes, will provide a solid support, fiber-optic properties similar to those of fibers utilized framework for the application of recently developed molecular in the telecommunications industry (5, 385). Importantly from tools for ecophysiological analyses of uncultivated microbes (2, a technical point of view, spicules are formed at ambient tem- 445, 449). Moreover, future combinations of (hypothesis-gen- peratures, and the study of this process may allow some of the erating) metagenomic data with such tools should be particu- inherent problems associated with the commercial (high-tem- larly profitable, especially for enigmatic microbes such as the perature) manufacture of optical fibers to be overcome (385). “Poribacteria.” Another area warranting further attention is Further insights into how organisms synthesize such sophisti- that of the molecular and biochemical bases of sponge-microbe cated biological materials were gained from electron micros- interactions. Recent work on the innate immune responses of copy observations of spicule organization in the same sponge sponges has already provided many important insights, but (6). The spicules of E. aspergillum provide great structural much remains unknown about host-symbiont signaling and stability due to their intricate arrangement at several spatial regulation. levels, ranging from nanometers to centimeters, perhaps in- Although many aspects of sponge-microbe associations are spiring the future development of new materials by humans interesting and important from a basic research perspective, (6). As mentioned earlier, sponges (e.g., the hexactinellid we acknowledge that it is largely biotechnological interest that sponge Scolymastra joubini) (51) may derive at least some of will sustain this field into the future. It is thus fitting to end their silica from diatoms, while in turn, spicules may extend the our review with this topic. The biotechnological potential of range of photosynthetic symbionts within sponge tissue via sponge-microbe associations has been widely (and justifiably) their conduction of light (115). Ongoing work on sponge skel- lauded, yet the transition from initial compound discovery to etogenesis (e.g., see references 245, 357, 418, and 490) should large-scale commercial production remains difficult. Chemical provide fascinating insights into its biological and, potentially, synthesis of sponge-derived compounds or their simpler deriv- commercial implications. atives offers the most reliable option for sustainable, long-term drug supply if the said compounds can be produced cost-effec- CONCLUSIONS AND FUTURE DIRECTIONS tively. Emerging technologies such as metagenomics and high- throughput microbial cultivation approaches offer exciting po- As indicated at the beginning of this article, even the signif- tential for accessing those compounds which are produced by icant recent advances in our understanding of sponge-micro- microorganisms. Current problems for metagenomics due to organism associations have not closed numerous gaping holes the complexity of microbial communities in sponges may be in our knowledge of these systems. It is startling how little is overcome in the future by single-cell genomic approaches known about many fundamental aspects of sponge symbiont (such as multiple displacement amplification) applied to spe- biology, particularly in the areas of symbiont metabolism and cific cell types which have been separated by, e.g., FISH and evolution. On the other hand, the ever-increasing research flow cytometry (e.g., see reference 183). The use of sponge interest in this topic (Fig. 1) promises a bright future for the larvae as starting material for metagenomic studies of drug- field. To close, we offer our thoughts on where some of this producing microbes has also been suggested, as these are most research attention could best be directed. likely to represent the true symbionts and, perhaps, may be Detailed studies of symbiont transmission and sponge-mi- simpler communities (279). The demonstrated complexity of crobe coevolution will greatly facilitate our understanding of vertically transmitted communities (reviewed in this article) the evolution of these systems. Improved host phylogenies, suggests, however, that even this will be challenging. Combi- from species to class levels, will be critical in this regard, and natorial biosynthesis, as exemplified by an elegant recent study recent efforts in this direction are highly encouraging. “Hunt- of Prochloron sp. symbionts of ascidians (82), provides fur- ing” for apparently sponge-specific microorganisms in other, ther potential for exploiting the chemical novelties present in nonsponge habitats (e.g., seawater and other filter-feeding in- sponge-microbe associations. Looking to the future, it is clear vertebrates) will also be important. However, merely proving that even greater integration among microbiologists, chemists, the presence of a sponge-specific microbe outside a sponge geneticists, zoologists, and aquaculture experts will be crucial host is not sufficient to disprove its sponge-specific nature. For in order to wring the most (data) out of sponges and their example, a sponge damaged by a predator or storm may dis- microbial partners. integrate and spread its microbial inhabitants into the water column. It is thus necessary to prove the activity of such mi- ACKNOWLEDGMENTS croorganisms outside the sponge host, using methods which We gratefully acknowledge the constructive comments on various link microbial identity with function (e.g., FISH-microautora- sections of the manuscript provided by Rocky de Nys, Friederike diography or stable isotope probing). This will be no small feat Hoffmann, Peter Schupp, Bob Thacker, and Torsten Thomas. The if the organism is extremely rare in the ocean, as will almost ideas expressed in this article also benefitted from discussions and/or collaborations with Rocky de Nys, Piers Ettinger-Epstein, Ute certainly be the case. Hentschel, Matthias Horn, Staffan Kjelleberg, Peter Schupp, Peter The function and physiology of sponge-associated microbes Steinberg, and Steve Whalan. We thank Ute Hentschel for providing are increasingly important research topics, reflecting our cur- samples of Plakortis sp. and Agelas dilatata and Megan Huggett and

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Sharon Longford for collection of Antho chartacea. Ivica Letunic and activity and surface bacterial film in marine sponges. J. Exp. Mar. Biol. Peer Bork are thanked for their help with the phylogenetic tree dis- Ecol. 179:195–205. played in Fig. 4, while we are indebted to Rocky de Nys for his 20. Becerro, M. A., R. W. Thacker, X. Turon, M. J. Uriz, and V. J. Paul. 2003. compilation of the chemical structures comprising Fig. 21 to 23. Jo¨rg Biogeography of sponge chemical ecology: comparisons of tropical and Ott kindly facilitated access to some of the sponge photographs in temperate defenses. Oecologia 135:91–101. 21. Belarbi, E., M. R. Dominguez, M. C. C. Garcia, A. C. Gomez, F. G. Camacho, Fig. 3. We thank Kayley Usher, Russell Hill, and Nicole Webster for and E. M. Grima. 2003. Cultivation of explants of the marine sponge their permission to reproduce published figures. Crambe crambe in closed systems. Biomol. Eng. 20:333–337. The research of M.W.T. was funded in part by a grant from the 22. Belarbi, E. H., A. C. Gomez, Y. Chisti, F. G. Camacho, and E. M. Grima. German Federal Ministry for Education and Research (Biolog II) to 2003. Producing drugs from marine sponges. Biotechnol. Adv. 21:585–598. M.W. D.S. received a DOC-FORTE stipend from the Austrian Acad- 23. Bell, A. H., P. R. Bergquist, and C. N. Battershill. 1999. Feeding biology of emy of Sciences. M.W. thanks the University of Vienna for financial Polymastia croceus. Mem. Qld. Mus. 44:51–56. support in the framework of the faculty focus “Symbiosis” and the 24. Bergmann, W., and R. J. Feeney. 1951. Contributions to the study of marine university research focus “Symbiosis Research and Molecular Princi- products. 32. The nucleosides of sponges. J. Org. Chem. 16:981–987. 25. Bergmann, W., and R. J. Feeney. 1950. The isolation of a new thymine ples of Recognition.” pentoside from sponges. J. Am. Chem. Soc. 72:2809–2810. 26. Bergquist, P. R. 1978. 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179

1 Supplemental Material for Taylor et al., “Sponge-associated microorganisms: evolution,

2 ecology and biotechnological potential”

3

4

5 Supplemental Figures

6

7 Suppl. Fig. 1 16S rRNA-based phylogeny of sponge-associated Bacteroidetes. Details are as

8 provided for Figs. 5-7 in the article.

9 Suppl. Fig. 2 16S rRNA-based phylogeny of sponge-associated Firmicutes. Details are as

10 provided for Figs. 5-7 in the article.

11 Suppl. Fig. 3 16S rRNA-based phylogeny of sponge-associated Actinobacteria (excluding

12 Family Acidimicrobiaceae – shown in Fig. 13). Details are as provided for Figs. 5-7 in the

13 article.

14 Suppl. Fig. 4 16S rRNA-based phylogeny of sponge-associated Alphaproteobacteria. Other

15 members of this class are shown in Fig. 14 and Suppl. Fig. 5. Details are as provided for Figs.

16 5-7 in the article.

17 Suppl. Fig. 5 16S rRNA-based phylogeny of sponge-associated Alphaproteobacteria. Other

18 members of this class are shown in Fig. 14 and Suppl. Fig. 4. Details are as provided for Figs.

19 5-7 in the article.

20 Suppl. Fig. 6 16S rRNA-based phylogeny of sponge-associated Gammaproteobacteria.

21 Other members of this class are shown in Suppl. Figs. 7 and 8. Details are as provided for

22 Figs. 5-7 in the article.

23 Suppl. Fig. 7 16S rRNA-based phylogeny of sponge-associated Gammaproteobacteria.

24 Other members of this class are shown in Suppl. Figs. 6 and 8. Details are as provided for

25 Figs. 5-7 in the article.

181 1 Suppl. Fig. 8 16S rRNA-based phylogeny of sponge-associated Gammaproteobacteria.

2 Other members of this class are shown in Suppl. Figs. 6 and 7. Details are as provided for

3 Figs. 5-7 in the article.

4 Suppl. Fig. 9 16S rRNA-based phylogeny of sponge-associated Betaproteobacteria. Details

5 are as provided for Figs. 5-7 in the article.

6

7

8 DNA extraction, PCR amplification, and cloning of 16S rRNA from three marine sponges

9

10 The sponges Agelas dilatata and Plakortis sp. were collected from Little San Salvador Island,

11 Bahamas, and kindly provided to us by Dr Ute Hentschel (University of Würzburg). Antho

12 chartacea was collected from Bare Island, Sydney, Australia by Sharon Longford and Megan

13 Huggett, whose efforts we gratefully acknowledge. Collections were performed under as

14 sterile conditions as possible, and samples were frozen prior to later freeze-drying.

15 We extracted DNA from 5 mg of freeze-dried tissue, from each of three individuals

16 per sponge species, using the UltraClean Soil DNA Isolation Kit (MoBio), according to the

17 manufacturer’s instructions. PCR amplification was performed in a BioRad iCycler, using the

18 following primers:

19 616V (5’- AGA GTT TGA TYM TGG CTC -3’) and 1492R (5’- GGY TAC CTT GTT ACG

20 ACT T -3’) for Bacteria (616V and 1390R (5’- GAC GGG CGG TGT GTA CAA -3’) were

21 used for amplifications with A. chartacea)

22 ARCH21F (5’- TTC CGG TTG ATC CYG CCG GA -3’) and 1492Rdeg (5’- GGY YAC

23 CTT GTT ACG ACT T -3’) for Archaea

24 POR389F (5’- ACG ATG CGA CGC CGC GTG -3’) and POR1130R (5’- GGC TCG TCA

25 CCA GCG GTC -3’) for “Poribacteria”

182 1 Cycling conditions were: 94°C for 3 min; 30 cycles of 94°C for 40 s, 52°C for 40 s, 72°C for

2 1 min 30 s; 72°C for 10 min.

3 Correct-sized amplicons were excised from a low-melting point agarose gel, then

4 cloned using the TOPO-TA Cloning Kit (Invitrogen), according to the manufacturer’s

5 instructions. Clones containing an insert of the correct length were selected randomly for

6 sequencing. The obtained sequences are available in Genbank under accession numbers

7 EF076059 to EF076242.

8

9

10 Construction of an ARB database comprising all publicly available sponge-derived 16S

11 rRNA sequences

12

13 We imported into ARB all sequences which were retrieved from Genbank on 1 March 2006

14 using the search string: (sponge* or porifera*) and (16S* or ssu* or rRNA*). We then

15 removed all extraneous sequences, including 18S rRNA sequences derived from the sponge

16 genome, and 16S rRNA sequences which contained ‘sponge’ somewhere in the Genbank

17 record but were in fact e.g. seawater-derived. After removing 26 sequences due to

18 insufficient quality for reliable alignment, and adding a further 184 16S rRNA sequences

19 from the sponges Agelas dilatata, Antho chartacea and Plakortis sp. (see below), we were left

20 with a total of 1683 sponge-derived 16S rRNA sequences and 11 18S rRNA sequences from

21 eukaryotic microbes in sponges. The annotated ARB database and alignment, with these

22 1694 sponge-derived 16S and 18S rRNA sequences, is available from the authors upon

23 request.

24

25

26 Phylogenetic analyses of sponge-derived 16S rRNA sequences

183 1

2 Preliminary analyses were performed to ascertain which sponge-derived sequences fell into

3 putative monophyletic, sponge-specific clusters. Representatives of every such cluster were

4 then screened against both the regular and environmental NCBI databases using the BLAST

5 function. The several most similar sequences to those in putative sponge-specific clusters

6 were then imported into ARB and aligned as described above. These and other related

7 sequences were used as reference sequences for subsequent phylogenetic analyses.

8 The majority of phylogenetic analyses were performed in the ARB programme

9 package. Maximum likelihood, maximum parsimony and neighbour-joining trees were

10 constructed on long (≥1000 nt) sequences using the programs AxML, DNAPARS and

11 Neighbor Joining, respectively. In three cases (Actinobacteria, Alphaproteobacteria and

12 Beta/Gammaproteobacteria) the number of sequences was too high to perform the maximum

13 likelihood analyses in ARB, so the program RAxML-VI-HPC was used (2). We compared

14 the topology of smaller trees obtained with AxML (implemented in ARB) and RAxML-VI-

15 HPC, and the results were highly consistent. Short sequences (>1000 nt) were subsequently

16 added to trees using the Parsimony Interactive tool in ARB, with no apparent change in tree

17 topology. Bootstrap analyses (100 resamplings) were performed in ARB using the

18 DNAPARS program, except for the Actinobacteria, Alphaproteobacteria and

19 Beta/Gammaproteobacteria trees, for which neighbour-joining bootstraps were calculated in

20 PAUP* 4.0b 10 (PPC). Positional conservation filters of 50% were applied for each

21 respective phylum, with outgroups consisting of sequences representing several other

22 bacterial phyla.

23 Maximum likelihood trees are displayed in the article, with sponge-specific clusters

24 only marked if they occurred consistently with all three treeing methods (some clusters are an

25 exception, occurring only in two methods – these are marked accordingly in the relevant

184 1 figures). The circular tree displayed in Fig. 4 was calculated in RAxML-VI-HPC, then

2 refined using the tools available at itol.embl.de (1).

3

4

5 References

6

7 1. Letunic, I., and P. Bork. 2006. Interactive Tree Of Life (iTOL): an online tool for

8 phylogenetic tree display and annotation. Bioinformatics publ. online.

9 2. Stamatakis, A. 2006. RAxML-VI-HPC: maximum likelihood-based phylogenetic

10 analyses with thousands of taxa and mixed models. Bioinformatics 22:2688-2690.

11

185 Suppl. FIG. 1 - Taylor et al.

Dysidea avara clone D.01, DQ274132 Dysidea avara clone D.29, DQ274137 Dysidea avara clone D.03, DQ274133 Dysidea avara clone D.128, DQ274150 Bizionia paragorgiae, AY651070 Dysidea avara clone D.79, DQ274144 Dysidea avara clone D.91, DQ274146 Dysidea avara DGGE band C4, AY947742 Dysidea avara DGGE band C9, AY947747 Arctic isolate 4-3, AF513398 Antarctic isolate IC164, AF001372 Phakellia ventilabrum isolate R8-Ret-T53-23g, AM117936 Psychroserpens burtonensis, AY771723 Craniella australiensis DGGE band D9, AY947737 Phakellia ventilabrum clone R8-Ret-T12_11d, AM117935 Coelosphaeridae isolate L303, AY371412 Myxilla incrustans clone UST050418-085, DQ202393 Antarctic isolate IC148, AF001373 Antarctic isolate IC159, AF001371 Halichondria panicea isolate Ex5, AY94836 Halichondria panicea isolate Ex9, AY948369 bloom isolate D21, AY030100 marine clone KMM3907, AY521223 Lissodendoryx isodictyalis isolate UST030701-295, AY848823 Suberites clone S2-1, AJ810791 Suberites clone S2-4, AJ810793 Suberites clone S2-12, AJ810796 Psychroserpens burtonensis, U62912 Arctic sea clone ARK10144, AF468421 Halichondria panicea isolate 698, Y17132 Gelidibacter algens, U62914 Gelidibacter mesophilus, AJ344134 coral isolate 'culture 8', DQ312231 sponge (5 sponges + sea urchin) isolate E966, AY368517 salt marsh isolate BSD S2 05, AY259512 Ancorinidae isolate A973, AY371406 marine bacterioplankton isolate GMD16C04, AY162110 Kirkpatrickia varialosa clone K102, AY321386 Aequorivita antarctica, AY027802 uranium mine tailing clone GR-Sh1-5, AJ296578 Vitellibacter vladivostokensis, AB071382 Pseudoceratinidae isolate K413, AY362005 estuary clone CF58, AY274858 Siphonidiidae isolate R564, AY367761 aquatile, M62797 industrial water isolate 3A5, AF368756 Petrosiidae isolate L979, AF486815 sponge (sponge + sediment) isolate M775, AY517542 Muricauda aquimarina, AY445076 Muricauda flavescens, AY445073 Rhabdastrella globostellata isolate 335, AY372916 marine alga isolate MBIC04669, AB073576 sponge isolate T-424, AB073584 Zobellia uliginosa, M62799 Pseudoceratina clavata isolate HPC1, AY372917 Arenibacter latericius, AF052742 Dysidea avara clone D.39, DQ274139 Flexibacter aggregans, AB078039 Cellulophaga pacifica, AB100842 Cellulophaga baltica, AJ005972 Halichondria panicea clone HOD12, Z88574, Cellulophaga lytica, AB032512 Craniella australiensis DGGE band D1, AY947729 Halichondria panicea isolate Ex26, AY948371 Halichondria panicea isolate Zo26, AY948381 Cytophaga latercula, M58769 Psychroflexus torquis, U85881 Salegentibacter salegens, M92279 Lissodendoryx isodictyalis isolate UST030701-156, DQ064789 sponge (4 sponges + sediment) isolate J879, AY370005 forest soil clone UC1, AY080916 ML, MP Halichondria panicea clone HRV16, Z88588 Halichondria panicea clone HRV11, Z88587 Axinella corrugata clone OS001, AY542747 Arctic bacterioplankton clone Arctic96B-24, AF354616 Mycale adherens isolate UST010723-003, AY241398 Tenacibaculum mesophilum, AB032503 sponge isolate T-551, AB073586 Tenacibaculum skagerrakense, AF469612 Flexibacter aurantiacus, AB078044 Axinellidae isolate H406, AY367750 Halichondria panicea isolate Zo6, AY948374 Halichondria panicea isolate 697, Y17133 Tenacibaculum maritimum, M64629 Halichondria panicea isolate Zo9, AY948376 Polaribacter irgensii, AY167325 Kirkpatrickia varialosa clone K21, AY321390 Polaribacter franzmannii, U14586 marine bacterioplankton clone ZD0255, AJ400343 Antarctic isolate MGP-9, AF530135 Dysidea avara DGGE band C8, AY947746 diatom detritus clone G1B2, AY285943 Mycale acerata clone M11, AY321405 Roseivirga seohaensis, AY739663 Roseivirga echinicomitans, AY753206 sponge isolate R966, AY371413 dinoflagellate-associated isolate DG1129, AY258133 Tedania ignis isolate UST030701-087, DQ080995 Tedania ignis isolate UST030701-084, DQ080996 sponge (2 sponges) isolate K439, AY367763 marine isolate MBIC01599 , AB086624 sponge (2 sponges) isolate K383, AF489285 coral isolate 'culture 9', DQ312232 Algoriphagus antarcticus, AJ577141 Flexibacter tractuosus, M58789 "Microscilla sericea", AB078081 "Microscilla furvescens", AB078079 Mycale laxissima clone MLLC14, DQ098842 Mycale laxissima clone MLLC12, DQ098840 Mycale laxissima clone MLLC15, DQ098843 Mycale laxissima clone MLLC11, DQ098839 marine bacterioplankton clone AEGEAN_150, AF406540 Sphaerotylus antarcticus clone S27, AY321414 hydrothermal vent worm clone P. palm A 21, AJ441237 Flexibacter litoralis, M58784 Flexibacter polymorphus, M58786 Flexibacter filiformis, AB078049 Flavobacterium ferrugineum, M62798 Cytophaga hutchinsonii, M58768 sponge isolate I-482, AB073593 Pseudoceratinidae isolate K429, AY367760 Flavobacterium mizutaii, AJ438175 Sphingobacterium multivorum, AB100739 "Sphingobacterium comitans", X91814 Bacteroides fragilis, X83938 Bacteroides thetaiotaomicron, L16489 Bacteroides acidifaciens, AB021164 Prevotella denticola, AY323524 Prevotella ruminicola, AF218618 Prevotella bryantii, AF396925 Bacteroides splanchnicus, L16496 Cytophaga fermentans, M58766 human intestine clone Z652, AY979341 Antho chartacea clone AnCha229f, EF076239 Bacteroides putredinis, L16497 Aplysina cavernicola DGGE band 1, AY180076 methanogenic consortium clone UASB_TL54, AF254396 hydrothermal vent clone AT-co1, AY225660 Cymbastela concentrica clone Cc180, AY942779 Kirkpatrickia varialosa clone K126, AY321388 Kirkpatrickia varialosa clone K6, AY321391 Sargasso sea metagenome clone IBEA_CTG_2050753, AACY01107683 marine bacterioplankton clone Arctic97A-17, AF354617 waste-gas biolfilter clone Blfcii9, AJ318122 Arctic clone ARKIA-105, AF468278 estuary clone 1A5, AY274868 Halichondria panicea clone HOS19, Z88577 Latrunculia apicalis clone L33, AY321399 soda lake clone ML617.5J-5, AF507867 Flexibacter aggregans, M58791 Dysidea avara DGGE band C7, AY947745 deep-sea sediment clone D66, AY375149 Rhopaloeides odorabile clone R63, AF333538 Aplysina aerophoba clone TK44, AJ347046 dinoflagellate-associated isolate ISR03, Y14143 Dysidea avara clone D.98, DQ274147 Theonella swinhoei clone RSWS16, AF434944 Theonella swinhoei clone JAWS2, AF434975 Theonella swinhoei clone RSWS18, AF434946 Discodermia dissoluta clone Dd-spU-17, AY897094 Chondrilla nucula clone Cnuc9, DQ079050 marine sediment clone S.Ionian-E03, AY534039 soil clone 9, AY493957 Salinibacter ruber, AY531231 deep-sea sediment clone MBAE20, AJ567581 Rhodothermus marinus, AF217494 deep-sea sediment clone BD2-15, AB015543

0.10 186 Pseudoceratina clavata isolate PC8, AY372919 Suppl. FIG. 2 - Taylor et al. Rhabdastrella globostellata isolate JS3, AY372923 Pseudoceratina clavata clone M108, DQ227667 Rhabdastrella globostellata isolate JS15, AY372921 Rhabdastrella globostellata isolate JS12, AY372920 sponge (sponge + sediment) isolate M608, AY364582 Lissodendoryx isodictyalis isolate SP8, AY780434 Bacillus firmus, D78314 Stelletta tenui clone, DQ274112 Bacillus firmus, AY833571 Pseudoceratina clavata isolate M13, DQ227670 Pseudoceratina clavata clone DE06, DQ227669 Pseudoceratina clavata clone F22, DQ227668 Siphonidiidae isolate L795, AY364586 Desulfotomaculum thermobenzoicum, Y11574 Geobacillus thermoleovorans, M77488 Bacillus methanolicus, X64465 Bacillus jeotgali, AF221061 sponge (3 sponges) isolate, AY364593 Exiguobacterium acetylicum, X70313, Mycale laxissima clone MLLC03, DQ098831 Pseudoceratina clavata clone M110, DQ227671 Bacillus benzoevorans, D78311 sponge (9 sponges + sediment) isolate P313, AY364581 Bacillus circulans, X60613 sponge (10 sponges) isolate B940, AY364590 Aplysina aerophoba isolate, AF218239 sponge (3 sponges) isolate E051, AY364587 Bacillus niacini, AB021194 soil clone 335-1, AF423249 sponge symbiont isolate HJ302, DQ167240 glacial ice clone SB100-9-5-1, AF479372 Amphimedon compressa isolate SP6, AY780433 Bacillus cohnii, X76437 Bacillus azotoformans, X60609 sponge (4 sponges & sediment) isolate J383, AY371403 Candidaspongia flabellata isolate AB004, AF369643 Bacillus fastidiosus, X60615 Hymeniacidon perleve isolate NJ632, AY621379 Pseudoceratina clavata isolate PC1, AY372926 maize rhizosphere isolate M4, AY376876 Bacillus megaterium, X60629 Pachastrellidae isolate J357, AY371404 Halichondria rugosa clone B14, DQ277984 Bacillus flexus, AB021185 Halichondria rugosa clone B02, DQ277980 Rhopaloeides odorabile clone NW-Sp3BO, AF333557 sponge (5 sponges) isolate D727, AY364589 Bacillus thuringiensis, AF290545 Bacillus anthracis, AY138366 Bacillus cereus, AY319260 sponge (4 sponges + sea cucumber + urchin) isolate F804, AY371400 Stelletta tenui clone A77, DQ274119 Stelletta tenui clone A71, DQ274118 Halichondria rugosa clone B31(2005), DQ277987 Agelas conifera isolate SP5, AY780435 Bacillus aquimaris, DQ432010 marine isolate JC1076, AF221855 Pseudoceratina clavata clone SW11, DQ227666 Geodiidae isolate D516, AY371689 oil-contaminated marine sediment isolate MN-003, AF355627 Psammplysilla purpurea clone PS2, AB196350 Psammplysilla purpurea clone PS79, AB196353 salt plain clone MO12, AY553105 Bacillus licheniformis, X68416 Mycale adherens isolate UST010723-004, AY241399 Rhopaloeides odorabile clone NW-Sp3Y, AF333558 Rhopaloeides odorabile clone NW-Sp3AS, AF333556 Halichondria rugosa clone B04, DQ277981 Halichondria rugosa clone B05, DQ277982 Bacillus mojavensis, AB021191 Halichondria rugosa clone B18, DQ277986 Aplysina aerophoba isolate A202, AF447805 glacial ice isolate, AF479349 Aplysina aerophoba isolate A190, AF447804 Aplysina aerophoba isolate A184, AF447803 Bacillus subtilis, AB042061 Psammplysilla purpurea clone PS9, AB196351 dry smoked sausages isolate Te68R, AY587832 Aplysina aerophoba isolate A586, AF447806 Bacillus pumilus, AY456263 Axinellidae isolate H762, AY364588 Halichondria rugosa clone B61(2005), DQ277988 Halichondria rugosa clone B15, DQ277985 Stelletta tenui clone 66, DQ236258 Stelletta tenui clone 68, DQ236259 Stelletta tenui clone 27, DQ236255 Sporosarcina pasteurii, X60631 Halichondria rugosa clone H.241, DQ274127 Halichondria rugosa clone H.323, DQ274130 Sporosarcina globispora, X54967 Sporosarcina psychrophila, X54968 Bacillus fusiformis, L14013 Pseudoceratina clavata´isolate PC2, AY372918 Kurthia gibsonii, X70320 Planococcus citreus, X62172 Azoricidae isolate H184, AY368541 Craniella australiensis DGGE band D6, AY947734 Stelletta tenui clone A45, DQ274116 Stelletta tenui clone A64, DQ274117 Bacillus sporothermodurans, U49078 Bacillus badius, X77790 Axinellidae isolate H761, AY364583 Bacillus gibsonii, X76446 Plakortis simplex isolate P203, AJ880003 Bacillus clausii, X76440 Bacillus pseudalcaliphilus, X76449 Bacillus halodurans, AJ302709 Rhabdastrella globostellata isolate JS4, AY372924 Brevibacillus invocatus, AF378231 Pseudoceratina clavata isolate PC6, AY372925 Brevibacillus thermoruber, Z26921 Paenibacillus curdlanolyticus, D78466 Pachastrellidae isolate N290, AY371688 Aplysina aerophoba isolate, AF218243 marine sediment isolate CNJ904 PL04, DQ448797 sponge (2 sponges + sea cucumber) isolate H819, AY364591 Bacillus barbaricus, AJ422145 Siphonidiidae isolate L794, AY364584 estuarine isolate TP-H, AY536561 Rhabdastrella globostellata isolate JS13, AY372927 Halobacillus litoralis, X94558 Candidaspongia flabellata isolate AB007, AF369646 marine clone, AY351787 Amphibacillus xylanus, D82065 Psammplysilla purpurea clone PS11, AB196352 Virgibacillus pantothenticus, X60627 Haliclona sp. isolate HAL6, AJ849368. Lentibacillus salicampi, AY057394 Pseudoceratina clavata clone M118, DQ227673 Pseudoceratina clavata clone M115, DQ227672 Bacillus horti, D87035 Stelletta tenui clone A32, DQ274115 Staphylococcus epidermidis, D83363 Staphylococcus aureus, D83357 sponge (2 sponges) isolate G779, AY369994 sponge (4 sponges + sea cucumber) isolate J688, AY369995 Staphylococcus hominis, L37601 Craniella australiensis DGGE band D3, AY947731 sponge (14 sponges) isolate J318, AY369993 Staphylococcus haemolyticus, L37600 Mycale adherens isolate UST010620-004, AY241414 Homaxinella balfourensis clone H50, AY321382 Staphylococcus sciuri, AJ421446 Halichondria rugosa clone B118, DQ277990 Listeria innocua, X56152 Marinilactibacillus psychrotolerans, AB083412 sponge isolate M13-7, AB083411 sponge isolate M13-2, AB083406 sponge isolate M13-4, AB083408 sponge isolate M13-3, AB083407 sponge isolate M13-6, AB083410 sponge isolate M13-5, AB083409 Marinilactibacillus piezotolerans, AY485792 Lactobacillus plantarum, D79210 Enterococcus faecalis, AB012212 Streptococcus pneumoniae, AF003930 Latrunculia apicalis clone L27, AY321397 Peptostreptococcus anaerobius, L04168 Clostridium difficile, X73450 "Epulopiscium fishelsoni", AF067413 Halichondria okadai clone HOC39, AB054173 Mycoplasma hominis, M24473 Mycoplasma pneumoniae, M29061 Spiroplasma montanense, AY189307 Chondrilla nucula clone Cnuc40412, DQ079052 Mammoth Cave clone CCM16a, AY221049 Rhopaloeides odorabile clone R185, AF333542 Chondrilla nucula clone Cnuc11, DQ079053 Coprothermobacter platensis, Y08935 Coprothermobacter proteolyticus, X69335

0.10 187 Suppl. FIG. 3 - Taylor et al. Streptomyces fradiae, AB184069 sponge isolate N12, AY944250 sponge (6 sponges) isolate L732, AY370000 Hymeniacidon perlevis isolate HPA53 , DQ144228 Streptomyces albogriseolus, AJ494865 Streptomyces coelicolor, AL939124 Streptomyces violaceoruber, AF503494 sponge isolate R03, AY944262 Hymeniacidon perlevis isolate HPA5, DQ144226 Hymeniacidon perlevis isolate HPA55, DQ144229 Streptomyces tendae, D63873 sponge isolate H02, AY944256 sponge isolate C06, AY944259 Hymeniacidon perlevis isolate HPA136, DQ144217 Streptomyces aureofaciens, AY289116 Theonella swinhoei clone RSWS13, AF434942 Streptomyces speibonae, AF452714 Hymeniacidon perlevis isolate HPA160, DQ144220 Hymeniacidon perlevis isolate HPA1, DQ144212 Hymeniacidon perlevis isolate HPA50, DQ144227 Hymeniacidon perlevis isolate HPA125, DQ144214 sponge isolate N02, AY944251 Streptomyces albidoflavus, Z76676 Streptomyces coelicolor, Z76678 Streptomyces sampsonii, D63871 sponge isolate S01, AY944263 Axinella corrugata clone OS044, AY542790 Axinella corrugata clone OS033, AY542779 sponge isolate H01, AY944255 sponge isolate R02, AY944261 water-damaged building isolate VTT E-99-1336 (B329), AF429400 . Hymeniacidon perlevis isolate HPA72, DQ144231 Streptomyces intermedius, Z76686 Streptomyces bluensis, X79324 sponge isolate C14, AY944260 mangrove soil isolate 050642, AY529646 Phymaraphinidae isolate J379, AY371434 sponge isolate N13, AY944252 Hymeniacidon perlevis isolate HPA135, DQ144216 Hymeniacidon perlevis isolate HPA66, DQ144230 Hymeniacidon perlevis isolate HPB40, DQ144232 Hymeniacidon perlevis isolate HPA151, DQ144219 Streptomyces caviscabies, AF112160 sponge isolate S13, AY944265 Hymeniacidon perlevis isolate HPA131, DQ144215 Streptomyces flavogriseus, AJ494864 Streptomyces griseinus, AB184205 Hymeniacidon perlevis isolate s14, AY436315 Axinella corrugata clone OS028, AY542774 Axinella corrugata clone OS027, AY542773 Streptomyces galbus, X79852 Streptomyces bikiniensis, X79851 Pachastrellidae isolate D721, AY369999 Hymeniacidon perlevis isolate HPA124 , DQ144213 Streptomyces armeniacus, AB018092 Streptomyces rimosus subsp. rimosus, AB045883 Hymeniacidon perlevis isolate HPB43, DQ144233 Streptomyces thermolineatus, Z68097 sponge isolate H07, AY944257 Streptomyces megasporus, Z68100 sponge (5 sponges & other invertebrates) isolate K454, AF489290 sponge (sponge & coral) isolate L806, AY368523 Pachastrellidae isolate M057, AY371454 Scleritodermidae isolate L262, AY368524 rape rhizosphere isolate tsb086, AJ295457 Microbacterium barkeri, X77446 sponge (4 sponges, coral, polychaete) isolate E920, AY368525 Microbacterium luteolum, AB004718 Demospongiae isolate F873, AY367754 Microbacterium testaceum, X77445 Microbacterium arabinogalactanolyticum, AB004715 Pseudoceratina clavata isolate PC7, AY372902 sponge (2 sponges & coral) isolate D704, AY368516 Leucobacter komagatae, D17751 Halichondria panicea isolate Ex8, AY948368 Craniella australiensis clone D10, AY947738 Agromyces cerinus subsp. cerinus, X77448 Cryobacterium psychrophilum, D45058 sponge (8 sponges & coral) isolate K463, AY368526 Pachastrellidae isolate N321, AY371408 Curtobacterium flaccumfaciens, AJ312209 Janibacter brevis, AJ310085 Pseudoceratina clavata clone DE08, DQ227675 Rhabdastrella globostellata isolate HRG7, AY845398 sponge (7 sponges & coral) isolate K372, AY368515 Kytococcus sedentarius, X87755 Cellulosimicrobium variabile, AJ298873 Mycale laxissima clone MLLC06 , DQ098834 Halichondriidae isolate L560, AY367751 Halichondria panicea isolate Zo24, AY948380 hexane-degrading biofilm isolate MN 8.1d.1c, AJ313024 Aplysina aerophoba isolate, AF218240 Micrococcus antarcticus, AJ005932 Mycale adherens isolate UST010620-003 , AY241413 Micrococcus luteus, AF542073 Rhabdastrella globostellata isolate JS14 , AY372899 sponge (4 sponges) isolate J921, AY368527 Arthrobacter nicotianae, X80739 Arthrobacter agilis, X80748 Aplysina aerophoba isolate, AF218242 Candidaspongia flabellata isolate AB006, AF369645 Kocuria rhizophila, Y16264 Mycale adherens isolate UST010620-002 , AY241412 Pseudoceratina clavata isolate, AY372901 Mycale adherens isolate UST010620-001 , AY241411 Kocuria kristinae, X80749 sponge (13 sponges & coral) isolate J935, AY364598 Brevibacterium linens, X77451 Siphonidiidae isolate R603, AY367748 Corynebacterium nigricans, AF220220 Corynebacterium jeikeium, U87823 sponge (2 sponges) isolate N138, AY367749 Corynebacterium amycolatum, X82057 sponge (4 sponges) isolate E241, AY367752 Dietzia maris, X81920 sponge (4 sponges & sea urchin) isolate J970, AY367753

various unidentified sponges (3) ML, MP Tsukamurella pulmonis, X92981 Tsukamurella spumae, AY238513 sponge isolate K362, AF502958 Phloeodictyidae isolate J313, AY368530 Mycobacterium elephantis, AJ010747 Mycobacterium peregrinum, AF058712 . Halichondria bowerbanki isolate ATCC 35087, AF480589 Halichondria bowerbanki isolate Mycobacterium poriferae, AF547959 Halichondriidae isolate J855, AY368507 Hymeniacidon perlevis isolate HPA140 , DQ144218 Rhodococcus marinonascens, X80617 Rhodococcus koreensis, AF124342 Rhodococcus opacus, X80630 Axinella corrugata, Demospongiae, Pseudoceratina clavata (3) Hymeniacidon perlevis isolate HPA180 , DQ144223 Nocardia ignorata, AJ303008 sponge isolate N28, AY944254 Nocardia asteroides, X84850 Nocardia abscessus, AF218292 Nocardia brasiliensis, X80591 Hexactinellida isolate F867, AY368506 terrae, X53202 Gordonia hirsuta, X93485 Rhopaloeides odorabile clone, AF333554 Rhabdastrella globostellata isolate HRG6 , AY372898 Nocardia corynebacterioides, AY438619 Irciniidae isolate F786, AY368570 Rhodococcus ruber, X80625 sponge (3 sponges & sea urchin) isolate J012, AY362006 sponge (2 sponges) isolate J562, AY362007 Aeromicrobium fastidiosum, X76862 sponge (sponge, coral, sea urchin) isolate J561, AY368562 Pseudonocardia kongjuensis, AJ252833 Rhopaloeides odorabile clone NW-Sp3EI , AF333553 sponge isolate C07, AY944258 sponge isolate S07, AY944264 Pseudonocardia autotrophica, X54288 sponge isolate N16, AY944253 Hymeniacidon perlevis isolate HPA192 , DQ144225 Pseudonocardia alni, Y08535 Pseudonocardia compacta, X76959 sponge (4 sponges & coral) isolate B951, AY368561 Actinoalloteichus cyanogriseus, AB006178 Hymeniacidon perlevis isolate HPA177 , DQ144222 Pseudoceratina clavata (8) Salinispora tropica, AY040617 marine sediment isolate., AY040618 Pseudoceratina clavata isolate M101, AY562197 Pseudoceratina clavata isolate SW02, AY562203 Halichondriidae isolate L656, AY368529 Micromonospora matsumotoense, AF152109 Hymeniacidon perlevis isolate HPB6, DQ144234 Micromonospora aurantiaca, X92604 sponge (sponge & sea cucumber) isolate H775, AY368528) Axinella corrugata (4) Micromonospora echinospora subsp. echinospora, U58532 Axinella corrugata clone OS050, AY542796 Micromonospora rosaria, X92631 Axinella corrugata clone OS039, AY542785 Verrucosispora gifhornensis, Y15523 Axinella corrugata clone OS030, AY542776 Microsphaera multipartita, Y08541 Axinella corrugata clone OS008, AY542754 Nocardiopsis tropica, AF105971 Hymeniacidon perlevis isolate HPA173, DQ144221 Nocardiopsis aegyptia, AJ539401 Nocardiopsis xinjiangensis, AF251709 Nocardiopsis dassonvillei subsp. albirubida, X97882 Phymaraphinidae isolate J380, AY368532 Axinella corrugata clone OS048, AY542794 Hymeniacidon perlevis isolate HPA19, DQ144224 Axinella corrugata clone OS036, AY542782 Axinella corrugata clone OS045, AY542791 Axinella corrugata clone OS041, AY542787 Axinella corrugata clone OS003, AY542749 Axinella corrugata clone OS012, AY542758 Spongilla lacustris (3) freshwater clone STH11-4, DQ316380 Spongilla lacustris clone SL30, AY480056 Spongilla lacustris clone SL17, AY466485 acidic hot spring clone NZ2, AJ575545 Spongilla lacustris clone SL48, AY466490 freshwater clone N3, AJ575530 Spongilla lacustris clone SL25, AY466486 Spongilla lacustris clone SS22, AY598786 Sporichthya polymorpha, AB025317 Halichondria panicea clone 8, AY948357 to Halichondria panicea clone 11, AY948359 outgroup

Acidimicrobiaceae [Fig. 13]

0.10 188 Suppl. FIG. 4 - Taylor et al.

Mycale adherens isolate UST010723-016, AY241410 sponge (7 sponges & bryozoan) isolate E923, AY369990 Siphonidiidae isolate L801, AY362016 Discodermia dissoluta clone Dd-spU-34, AY897099 Pseudoceratina clavata clone F14, DQ227660 Pseudoceratina clavata clone D29, DQ227659 sponge isolate MBIC1876, AB026194 demospongiae isolate N286, AY369984 demospongiae isolate N354, AY371430 Halichondria panicea isolate Ex11, AY948370 squid isolate SOGA8, AJ244786 squid isolate SOGA14, AJ244791 Ruegeria atlantica, D88526 sponge (many sponges, sediment, other invertebrates) isolate J355, AY362009 Silicibacter lacuscaerulensis, U77644 Suberites domuncula isolate SDC1, AY265806 marine bacterioplankton isolate BBAT2, AF365993 sponge clone MBIC1887, AB026492 dinoflagellate-associated isolate TM1040, AY332662 Silicibacter pomeroyi, AF098491 Homaxinella balfourensis clone H47, AY321430 Dysidea avara DGGE band C13, AY947751 Dysidea avara DGGE band C12, AY947750 hypersaline mat isolate GL3, AF344287 Scleritodermidae isolate P638, AY368522 DMSP-degrading marine isolate LFR, L15345 Demospongiae isolate J356, AY368512 hydrothermal vent isolate NF18, AF254107 marine bacterioplankton clone CHAB-I-5, AJ240910 Halichondria panicea clone HRV3# HpaAS1, Z88581 Callyspongia sp. DGGE band 7, AY258605 Cymbastela concentrica, Halichondria panicea, Stylinos sp. (+1 non-sponge sequence) (4) Sargasso Sea metagenome clone IBEA_CTG_2112418, AACY01072860 Sargasso Sea metagenome clone IBEA_CTG_2000148, AACY01008274 Lamellodysidea chlorea clone 310P14, AY845238 Lendenfeldia chondrodes clone 36P5, AY845239 Lendenfeldia chondrodes clone 36P16, AY845240 Lendenfeldia chondrodes, Phyllospongia papyracea (3) Lamellodysidea chlorea clone 310P5, AY845237 dinoflagellate-associated isolate DFL-16, AJ534213 Thorectidae isolate H265, AY371428 Roseovarius tolerans, Y11551 Phyllospongia papyracea clone 34P11, AY845242 Phyllospongia papyracea clone 34P35, AY845243 coral isolate 92, AY654831 sponge (3 sponges & coral) isolate L544, AY370007 marine aggregate isolate HP29w, AY239008 dinoflagellate-associated isolate DG1127, AY258099 Kirkpatrickia varialosa/Mycale acerata DGGE band J, AY320221 Latrunculia apicalis clone L24, AY321396 Homaxinella balfourensis clone H41, AY321381 Arctic isolate ARK10207, AF468373 Arctic isolate ARK9990, AY167339 Kirkpatrickia varialosa clone K106, AY321387 Geodiidae clone L992, AF489286 Sulfitobacter brevis, Y16425 Roseobacter litoralis, X78312 Halichondria panicea isolate Ex7, AY948367 Halichondria panicea isolate Zo7, AY948375 Roseobacter denitrificans, M96746 isolate, U37762 sponge (4 sponges & coral) isolate J484, AY369983 Octadecabacter arcticus, U73725 Coelosphaeridae isolate L35, AY370003 coral clone 128-19L, AF473915 sponge (4 sponges, sediment, other invertebrates) isolate J486, AY368574 lake clone K2-53B, AY345413 Black Sea isolate TH1, AJ133762 whale-fall metagenome clone AHAA3001.g1, AAFZ01011523 dinoflagellate-associated isolate DG1128, AY258100 Ruegeria gelatinovorans, D88523 sponge (5 sponges & sea cucumber) isolate J483, AY369980 Homaxinella balfourensis, Kirkpatrickia varialosa, Mycale acerata (8) marine sediment isolate EI1*, AF254105 sponge (2 sponges) isolate J504, AY369979 squid isolate J8W, AF026463 sponge symbiont isolate UST020129-036, AY241443 coastal marine isolate WHOI JT-08, AY349460 Ancorinidae isolate J48, AY369982 Rhopaloeides odorabile isolate NW43, AF384141 sponge (3 sponges & coral) isolate H264, AY369978 sponge (2 sponges) isolate H454, AY368572 Roseobacter gallaeciensis, AY881240 paper mill effluent isolate Y4I, AF388307 Mycale adherens isolate UST010723-007, AY241402 Mycale adherens isolate UST010723-008, AY241403 Ruegeria algicola, X78315 Roseobacter gallaeciensis, Y13244 Cymbastela concentrica DGGE band 8.3-CcNSW, AY569396 Kirkpatrickia varialosa/Mycale acerata DGGE band K, AY320222 Jannaschia helgolandensis, AJ438157 Latrunculia apicalis clone L41, AY321420 deep-sea sediment clone D46, AY375143 brittlestar clone, U63548 Halichondria panicea, Microciona (8) deep-sea sediment clone B61, AY375078 sponge (7 sponges & coral) isolate J987, AY368534 sponge (7 sponges & coral) isolate M039, AY368535 sponge (2 sponges) isolate J364, AY368536 Paracoccus yeeii, AY014179 Paracoccus aminovorans, D32240 Dysidea avara clone D.122, DQ274148 Dysidea avara clone D.125, DQ274149 Rhodobacter veldkampii, D16421 Dysidea avara clone D.132, DQ274151 Rhodobacter sphaeroides, X53854 Agelas dilatata, Chondrilla nucula (5) "Rhodovulum imhoffii", AM180953 Chondrilla nucula clone Cnuc16, DQ079032 Chondrilla nucula clone Cnuc4041, DQ079039 Rhodovulum euryhalinum, D16426 Rhodovulum strictum, D16419 Lamellodysidea herbacea clone, AY845236 Sargasso Sea metagenome clone IBEA_CTG_2012131, AACY01026023 Cymbastela concentrica DGGE band 9.1-CcNSW, AY569383 Cymbastela concentrica clone Cc118, AY942774 Cymbastela concentrica clone Cc070, AY942765 Hirschia baltica, X52909 Hyphomonas jannaschiana, AJ227814 Maricaulis maris, AJ007807 ML, MP Axinella corrugata, Chondrilla nucula, Halichondria okadai (+1 non-sponge sequence) (6) Suberites clone S1, AJ810784 Pelagibacter ubique, AF510191 Kirkpatrickia varialosa DGGE band G, AY320218 coral clone OTU 15, DQ312248 marine clone PWB3, AB106120 Cymbastela concentrica clone Cc155, AY942776 Chondrilla nucula clone Cnuc55, DQ079038 marine bacterioplankton clone AEGEAN_207, AF406520 Pseudovibrio spp. and relatives [Fig. 14]

Cymbastela concentrica clone Cc068, AY942764 Cymbastela concentrica to Suppl. Cymbastela concentrica clone Cc173, AY942778 from two locations Cymbastela concentrica DGGE band 8, AY258606 Fig. 5 Cymbastela concentrica DGGE band 3, AY258601 Cymbastela concentrica DGGE band 4.1-CcGBR, AY569375 dinoflagellate-associated isolate DG943, AY258089 sponge (Pachastrellidae & other invertebrates) isolate D701, AY368521 biowaste reactor isolate TUT1018, AB098586 Halichondria panicea clone HNSS21, Z88572 waste gas-degrading clone BIhii25, AJ318143 demospongiae isolate L519, AY371414 Fulvimarina pelagi, AY178860 Aurantimonas coralicida, AY065627 sponge (3 sponges & sea cucumber) isolate K416, AY368505 Mesorhizobium loti, D14514 Mesorhizobium mediterraneum, L38825 Defluvibacter lusatiensis, AJ132378 sponge (5 sponges & coral) isolate J211, AY367744 Rhizobium radiobacter, AJ389903 sponge (3 sponges) isolate F813, AY362010 Rhizobium giardinii, U86344 Rhizobium leguminosarum, AY509899 sponge (9 sponges) isolate E916, AY368533 Brucella melitensis, L26166 sponge isolate Rhodobium marinum, D30790 sponge isolate Rhodobium marinum NCIMB2201, D14434 dinoflagellate-associated isolate DG948, AY258091

0.10 189 Suppl. FIG. 5 - Taylor et al.

to Suppl. Fig. 4

Halichondria panicea clone sponge_clone3, AY948355 Halichondria panicea clone HNS6, Z88566 freshwater reservoir clone HTD9, AF418955 Mammoth Cave clone CCM12a, AY221043 Devosia riboflavina, D49423 Dysidea avara DGGE band C10, AY947748 Halichondriidae isolate F820, AY362018 Antarctic sediment bacterium MGP-83, AF530152 Halichondria panicea clone HNSS12, Z88570 Dysidea avara DGGE band C3, AY947741 Antarctic sediment bacterium MGP-68, AF530148 Antarctic lake isolate R-9221, AJ441010 Cymbastela concentrica DGGE band 10-CcNSW, AY569385 Halichondriidae isolate J560, AY371451 Sphingomonas suberifaciens, D13737 sponge (2 sponges & sediment) isolate J345, AY364594 sponge (2 sponges) isolate L538, AY369991 Sphingopyxis terrae, D13727 Stelletta tenui DGGE band A10, AY947762 Axinellidae isolate L534, AY369992 Sphingomonas koreensis, AF131296 Sphingomonas paucimobilis, U20776 Sphingomonas echinoides, AB021370 Haliclona sp. isolate HAL66h, AJ849370 Irciniidae isolate F752, AY367756 Erythrobacter citreus, AF118020 sponge (3 sponges & coral) isolate K384, AY367757 Erythrobacter flavus, AF500004 sponge (many sponges, sediment, other invertebrates) isolate F761, AY371410 Petrosiidae isolate G265, AY371411 Scleritodermidae isolate L259, AY367758 Erythrobacter litoralis, AB013354 Porphyrobacter neustonensis, L01785 Methylocystis echinoides, AJ458473 forest soil clone UP7, AY080914 Methylobacterium mesophilicum, D32225 ML, MP Antho chartacea, Axinella corrugata, Suberites (11) Rhopaloeides odorabile clone NW-Sp2A, AF333555 Brevundimonas vesicularis, AJ007801 sponge (2 sponges & other invertebrates) isolate F996, AY367745 Brevundimonas intermedia, AJ007802 sponge (3 sponges & sea cucumber) isolate F991, AY364600 Caulobacter vibrioides, AJ009957 insect larva clone D, AJ459874 Halichondria sp. DGGE band B7, AY947727 Halichondria sp. DGGE band B6, AY947726 Spongilla lacustris clone SS15, AY598784 Halichondriidae isolate J526, AY369981 Roseomonas cervicalis, AY150047 Spongilla lacustris clone SS42, AY598791 Azospirillum irakense, X79737 soil isolate Ellin329, AF498711

Halichondria okadai clones (27)

marine clone Surf1.34, DQ071157 Plakortis sp. clone PK021, EF076075 Halichondria panicea clone HRV35, Z88590 Sargasso Sea metagenome clone IBEA_CTG_1999031, AACY01099207 marine bacterioplankton clone MB11B10, AY033300 marine bacterioplankton clone MB12F11, AY033316 Mycale laxissima clone MLLC22, DQ098850 Mycale laxissima clone MLLC04, DQ098832 Halichondria panicea clone HRV2, Z88580 Halichondria panicea clone HRV7, Z88584 Halichondria panicea clone HRV6, Z88583 Halichondria panicea clone HRV39, Z88591 marine bacterioplankton clone OM38, U70679 Sargasso Sea metagenome clone IBEA_CTG_1958364, AACY01099937 Aplysina aerophoba, Theonella swinhoei (3) ML, MP Chondrilla nucula, Phyllospongia papyracea, Theonella swinhoei (+1 non-sponge sequence) (5)

Cymbastela concentrica, Phyllospongia papyracea, Plakortis sp. (+1 non-sponge sequence) (9) Aplysina aerophoba clone TK97, AJ347054 marine clone 9NBGBact_92, DQ070813 deep-sea sediment clone D108, AY375135 deep-sea sediment clone MBMPE38, AJ567552 Aplysina aerophoba, Discodermia dissoluta, Theonella swinhoei (5)

Pachastrellidae, Phyllospongia papyracea & 2 other sponges (5) deep-sea sediment clone A6, AY373415 oligochaete clone 1c, AJ890093 Aplysina aerophoba clone TK03, AJ347025 Discodermia dissoluta clone Dd-spT-C26, AY897109 Theonella swinhoei clone RSWS1, AF434963 deep-sea sediment clone B21, AY375070 Mammoth Cave clone CCU22, AY221078 Axinella verrucosa, Cymbastela concentrica, Halichondria panicea (3) hot spring clone PK324, AY555804 alpine soil clone P2-12H, AY192272 Spongilla lacustris clone SS21, AY598785 soil isolate Ellin346, AF498728 Spongilla lacustris clone SS31, AY598790 demospongiae isolate M914, AY371399 Rhodospirillum rubrum, M32020 Magnetospirillum magnetotacticum, Y10110 Spongilla lacustris clone SL21, AY466474 Spongilla lacustris clone SL53, AY466479 Spongilla lacustris clone SL24, AY466475 Spongilla lacustris clone SL38, AY466476 Spongilla lacustris clone SL51, AY466478 Spongilla lacustris clone SL40, AY466477 Spongilla lacustris clone SL43, AY466481 Spongilla lacustris clone SL44, AY466482 Spongilla lacustris clone SL33, AY466480 rumen clone RFN49, AB009205 "Odyssella thessalonicensis", AF069496 amoeba endosymbiont clone UWC36, AF069962 Cymbastela concentrica clone Cc045, AY942762 to Ehrlichia canis, M73221 , L36217 outgroup Holospora obtusa, X58198 Halichondria panicea clone HOS16, Z88576 Kirkpatrickia varialosa DGGE band F, AY320217

0.10 190 Suppl. FIG. 6 - Taylor et al. Phloeodictyidae isolate J312, AY371448 Theonellidae isolate E164, AY371449 marine isolate OC25, AB038026 marine isolate SW12, AY902205 Axinellidae isolate L536, AY370017 Vibrio lentus, AJ278881 Isops phlegraei isolate Gp-MBA-3_1.3, AJ849367 Vibrio splendidus, X74724 Axinellidae isolate H412, AY370014 Halichondria panicea isolate Zo2, AY948372 Halichondria panicea isolate Zo5, AY948373 coral isolate 12, AY654744 marine clone Hdeep-1, AB055796 Vibrio chagasii, AJ316199 sponge (7 sponges & other invertebrates) isolate J252, AY370018 marine clone 4c, AF293974 Vibrio pomeroyi, AJ491290 Suberites domuncula isolate SubDo-inf1, AF324884 Mycale adherens isolate UST010723-010, AY241404 Halichondria DGGE band B4, AY947724 sponge (5 sponges) isolate D725, AY370015 Listonella anguillarum, X16895 , X56582 Vibrio rumoiensis, AB013297 Pachastrellidae isolate J462, AY370011 Vibrio fischeri, X70640 Vibrio salmonicida, X70643 coral isolate 8, AY654822 Aplysina aerophoba isolate, AF218244 Vibrio halioticoli, AB000390 Vibrio superstes, AY155585 Mycale adherens isolate UST010723-002, AY241397 Latrunculia apicalis clone L19, AY321394 Mycale acerata clone M144, AY321409 Vibrio fortis, AJ514916 Vibrio sp. LMG 20547, AJ316202 Geodiidae isolate K883, AY371447 sponge (2 sponges) isolate J555, AY370012 Vibrio shilonii, AF007115 . Vibrio mediterranei, X74710 Vibrio orientalis, X74719 Vibrio furnissii, X74704 Callyspongia sp. isolate UST040911-030, DQ156534 diseased Rhopaloeides odorabile isolate NW4079, AF384142 Mycale adherens isolate UST010723-012, AY241406 oyster clone PP-145.98, AJ296157 Vibrio coralliilyticus, AJ440005 Rhabdastrella globostellata isolate HRG12, AY372930 Rhabdastrella globostellata isolate HRG15, AY372929 Vibrio nigripulchritudo, X74717 fish isolate 167, AF493803 Theonellidae isolate J684, AY370016 Vibrio brasiliensis, AJ316172 Mycale adherens isolate UST010723-001, AY241396 Vibrio natriegens, X74714 Candidaspongia flabellata isolate AB003, AF369642 , M59161 sponge (9 sponges) isolate J608, AY371446 Vibrio diabolicus, X99762 Vibrio proteolyticus, X56579 Vibrio campbellii, X56575 Vibrio harveyi, X56578 Callyspongia sp. isolate UST040911-028, DQ156530 Callyspongia sp. isolate SUT040911-031, DQ156535 deep-sea hydrothermal vent clone BB4, AF319768 sponge (many sponges & other invertebrates) isolate J821, AY370013 Mycale adherens isolate UST010723-015, AY241409 Photobacterium angustum, D25307 Photobacterium phosphoreum, D25310 sponge (4 sponges & sea cucumber) isolate J551, AY368538 Photobacterium profundum, D21226 sponge (4 sponges, sediment, other invertebrates) isolate J169, AY368513 Vibrio calviensis, AF118021 marine isolate EN276, AB038023 sponge (2 sponges & other invertebrates) isolate J725, AY368539 coral isolate 56, AY654789 sponge (2 sponges) isolate J246, AY368537 Photobacterium damselae, X74700 Spongionella nigra isolate A3B-64-2, AB193754 Ferrimonas balearica, X93021 Homaxinella balfourensis clone H30, AY321378 Sphaerotylus antarcticus clone S29, AY321418 "Pantoea endophytica", AF130958 Pantoea agglomerans, AY691545 Erwinia amylovora, Z96088 Antho chartacea clone AnCha112f, EF076230 Antho chartacea clone AnCha1_3, EF076196 Escherichia coli, X80723 , AE005674 , AE016834 , AF276988 Dysidea avara DGGE band C1, AY947739 Serratia liquefaciens, AB004752 , AY114159 , AY043385 Latrunculia apicalis clone L43, AY321422 Serratia rubidaea, AB004751 Buchnera aphidicola, AB064514 Providencia stuartii, AF008581 fly pathogen clone ME, U20277 Halichondria rugosa isolate B12, DQ277983 intestinal clone D22, AY374116 Halichondria rugosa clone H.11, DQ274120 Halichondria rugosa clone H.95, DQ274125 Craniella australiensis DGGE band D8, AY947736 Phakellia ventilabrum isolate Ph-Ret-19, AM117934 Rheinheimera pacifica, AB073132 Rheinheimera baltica, AJ002006 sponge (2 sponges) isolate R246, AY368567 Geodia baretti isolate Gbf-Ret-3, AM117933 Thiocapsa roseopersicina, Y12364 marine bacterioplankton clone TW-7, AY028202 . Sargasso sea metagenome clone IBEA_CTG_2158063, AACY01038944 Haliclona sp. isolate HAL15c, AJ849372 abalone gut clone DC1, AF280818 Halichondria panicea clone Zo12, AY948378 Pseudoalteromonas haloplanktis, X67024 Pseudoalteromonas nigrifaciens, X82146 deep-sea sediment isolate AC-144, AJ519790 Pseudoalteromonas tetraodonis, X82139 Pseudoalteromonas undina, X82140 marine bacterioplankton clone Tw-4, AY028199 . Aplysina aerophoba isolate, AF218245 microbial mat isolate EBD, DQ218321 Halichondria panicea isolate Ex1, AY948362 Pseudoalteromonas carrageenovora, X82136 Aplysina aerophoba isolate , AF218246 Pseudoalteromonas espejiana, X82143 sponge (3 sponges) isolate F92, AY368542 Arctic isolate ARCTIC-P16, AY573035 Pseudoalteromonas agarivorans, AJ417594 Isops phlegraei isolate Gp-MBA-2_5.3.1, AJ849373 deep-sea tubeworm clone TW-4, AB042419 Haliclona sp. isolate HAL34, AJ849371 sponge (2 sponges) isolate J21, AY368544 deep-sea hydrothermal vent clone EPR2, AY394863 submarine volcano clone JB10, AY345372 sponge (12 sponges) isolate H72, AY368545 deep-sea isolate ws17, AJ704790 Theonella swinhoei clone PAUC53, AF186438 Higginsia strigilata isolate J173, AY371427 deep-sea sediment isolate 3032, AM110986 Mycale adherens isolate UST010723-006, AY769918 marine isolate S9, U80834 Mycale adherens isolate UST010723-006, AY241401 Placospongiae isolate H274, AY371425 Pseudoalteromonas tunicata, Z25522 Fascaplysinopsis reticulata isolate Pseudoalteromonas maricaloris, AF144036 Hymeniacidon perleve isolate NJ6-3-1, AY621063 Pseudoalteromonas piscicida, AB090233 Pseudoalteromonas peptidolytica, AF007286 sponge (2 sponges) isolate M60, AY371426 marine clone RE2-11, AF539778 Pseudoalteromonas rubra, X82147 Mycale adherens isolate UST010620-005, AY241415 sponge (2 sponges) isolate B94, AY368543 Pseudoalteromonas luteoviolacea, X82144 Pseudoalteromonas aurantia, X82135 sponge (2 sponges) isolate P630, AY362019 marine bacterioplankton clone Tw-2, AY028197 . Dysidea avara clone D.135, DQ274152 Colwellia maris, AB002630 . Colwellia piezophila, AB094412 Suberites clone S2-8, AJ810794 Colwellia psychrerythrea, U85841 Halichondria panicea clone Ex3, AY948363 Suberites clone S2-9, AJ810795 marine isolate UMB3E, AF505723 Antarctic isolate ANT8279, AY167268 ML, MP Axinellidae, Coelosphaeridae, Sphaerotylus antarcticus (3) hydrothermal vent clone PVB_54, U15115 Thalassomonas viridans, AJ294748 Scleritodermidae isolate N362, AY368514 Idiomarina baltica, AJ440214 Idiomarina zobellii, AF052741 Halichondria panicea isolate Zo10, AY948377 Psychromonas profunda, AJ416756 Suberites clone S17, AJ810790 coral isolate 63, AY654801 brackish water clone PH-B21N, AF513950 smoked salmon clone 270, AJ310692 Haliclona sp. isolate HAL19, AJ849374 Ectyoplasia ferox isolate SP2, AY780443 Sargasso Sea metagenome clone IBEA_CTG_2017930, AACY01134203 marine clone CHAB-IV-19, AJ240914 Alteromonas macleodii, Y18228 marine isolate July88, AY082664 Callyspongia sp. isolate UST040911-043, DQ156529 Callyspongia sp. isolate UST030701-068, DQ156528 sponge (11 sponges) isolate J589, AY362020 sponge (5 sponges & sediment) isolate F49, AY368546 Callyspongia sp. isolate UST040911-003, DQ156533 Mycale adherens isolate UST010723-005, AY241400 Mycale adherens isolate UST010723-011, AY241405 marine clone SUR-ATT-18, AF114501 Mycale laxissima isolate SP4, AY780437 marine clone DCM-ATT-18, AF114500 deep-sea clone 400m-ATT-20, AF114502 dinoflagellate-associated clone ATAM407_18, AF359529 . marine clone CHAB-V-29, AJ240916 Petrosiidae isolate G287, AY368547 marine clone KT0812A, AF239705 Glaciecola pallidula, U85854 Glaciecola punicea, U85853 sponge (2 sponges) isolate N352, AY371690 Pseudoalteromonas sagamiensis, AB063324 Dysidea avara, Stelletta tenui (4) Oceanisphaera litoralis, AJ550470 "Oceanimonas denitrificans", DQ097665 to Suppl. Fig. 8 Oceanimonas doudoroffii, AB019390 demospongiae isolate K649, AY362008 salmonicida, X74681 Halichondria panicea, Ircinia dendroides, Mycale acerata, Thorectidae & other sponges (6) Sargasso Sea metagenome clone IBEA_CTG_2158520, AACY01118821 Shewanella putrefaciens, U91552 Sargasso Sea metagenome clone IBEA_CTG_2153346, AACY01101508 sponge (5 sponges) isolate H836, AY369988 Shewanella algae, U91547 deep-sea hydrothermal vent isolate A6.mk, AF319767 . Mycale adherens isolate UST010723-014, AY241408 marine sediment clone Belgica2005/10-130-32, DQ351772 sponge (5 sponges) isolate J327, AY369989 Shewanella hanedai, U91590 Shewanella woodyi, AF003549 . sponge (3 sponges & sea cucumber) isolate H411, AY369987 Shewanella pealeana, AF011335 Latrunculia apicalis clone L7, AY321401 sediment MGP-6, AF530133 sponge (3 sponges & seawater) DGGE band D, AY320215 . coral clone OTU 13, DQ312246 Moritella viscosa, AJ132226

0.10 191 Suppl. FIG. 7 - Taylor et al.

Betaproteobacteria [Suppl. Fig. 9] Dysidea avara clone D.46, DQ274140 Dysidea avara DGGE band C6, AY947744 Dysidea avara DGGE band C5, AY947743 Arctic clone Arctic96B-9, AF354596 Pachastrella sp. isolate Pseudomonas pachastrellae, AB125367 Pachastrella sp. isolate Pseudomonas pachastrellae, AB125366 marine sediment isolate MBIC2027, AB030085 hydrothermal vent isolate HY-95/2-1, AJ294342 sponge (3 sponges & sea cucumber) isolate E762, AY368557 Axinellidae isolate H741, AY368558 Pseudomonas fluorescens, Z76662 Pseudomonas chlororaphis, D86004 Desmoxyidae isolate J187, AY368550 Pseudomonas mosselii, AF072688 sponge (6 sponges & coral) isolate H757, AY368555 Pseudomonas fulva, D84015 sponges (2 sponges) isolate J192, AY368551 Pseudomonas putida, Z76667 Axinellidae isolate H756, AY368548 deep-sea sediment clone BD5-7, AB015564 . Pseudomonas aeruginosa, X06684 Pseudomonas alcaligenes, Z76653 saltmarsh clone LCP-79, AF286035 sponge (5 sponges) isolate K458, AF489288 sponge (5 sponges) isolate K433, AF489289 sponge (5 sponges) isolate H786, AY368554 Pseudomonas mendocina, Z76674 swine effluent isolate CHNDP38, DQ337540 sulfide-oxidizing isolate N9-1, AF393509 Stelletta tenui clone 6, DQ236251 Halichondria rugosa clone H.60, DQ274124 Halichondria rugosa clone H.59, DQ274123 Pseudomonas sp. HS5322, AY443041 Pseudomonas anguilliseptica, X99540 Scleritodermidae isolate P664, AY368549 sponge (3 sponges) isolate E763, AY368560 Pseudomonas balearica, U26418 Suberites domuncula isolate SubDo-sym, AF324886 sponge isolate UST040317-107, DQ096578 sponge isolate UST040317-067, DQ096579 Suberites domuncula isolate PB2, AF482707 Suberites domuncula isolate PB1, AF482708 coral isolate 94, AY654829 Pseudomonas sp. CJ11014, AF500211 Microbulbifer cystodytense, AJ620879 Pseudoceratina clavata isolate F25, DQ227664 Pseudoceratina clavata isolate D09, DQ227661 Pseudoceratina clavata isolate D10, DQ227663 Pseudoceratina clavata isolate F27, DQ227662 Candidaspongia flabellata isolate AB008, AF369647 sponge (4 sponges & sediment) isolate J451, AY368552, sponge (5 sponges & sediment) isolate N276, AY368556 coral isolate 83, AY654820 Microbulbifer hydrolyticus, U58338 Microbulbifer salipaludis, AF479688 Mycale adherens isolate UST010723-013, AY241407 Kirkpatrickia varialosa DGGE band H, AY320219 marine isolate HTCC2207, AY386335 Coelosphaeridae isolate L193, AY371439 Phyllospongia papyracea clone 34P38, AY845232 Cellvibrio vulgaris, AF448513 Ancorinidae isolate J505, AY371435 "Candidatus Endobugula glebosa", AY532642 Teredinibacter turnerae, AY028398

Plakortis sp., Rhopaloeides odorabile (9)

Agelas dilatata clone AD026, EF076137 Rhopaloeides odorabile clone R58, AF333539 Phyllospongia papyracea clone 31P6, AY845230 manure-contaminated water clone 266ds10, AY212716 marine isolate HTCC2089, AY386332 marine clone EF100-91A10, AY627370 marine clone KI89A, AB021704 Haliclona sp. isolate HAL40b, AM117932 Arctic clone ARKDMS-49, AF468250 Mycale acerata clone M129, AY321406 Sphaerotylus antarcticus clone S35, AY321417 marine clone KTc1119, AF235120 Halichondria panicea clone HRV1, Z88579

Agelas dilatata, Antho chartacea, Aplysina aerophoba, Discodermia dissoluta, Theonella swinhoei (8)

Nullarbor cave clone wb1_P19, AF317768 deep-sea sediment clone E77, AJ966605 hydrothermal sediment clone AT-s80, AY225635 hydrothermal sediment clone AT-s26, AY225634 Nitrosococcus halophilus, AF287298 Nitrosococcus oceani, M96395 Theonella swinhoei clone RSWS6, AF434961 to Suppl. Thialkalivibrio jannaschii, AF329083 Fig. 8 Nitrococcus mobilis, L35510 marine sediment clone Belagica 2005/10-140-24, DQ351796 Halichondria panicea clone HNSS31, Z88573 Sargasso Sea metagenome clone IBEA_CTG_2077477, AACY01036570 deep-sea sediment clone BD6-6, AB015576 . Rhopaloeides odorabile clone R93, AF333534 deep-sea sediment clone A126, AY373398 Cymbastela concentrica clone Cc190, AY942781 deep-sea sediment clone NKB5, AB013257 . , D89791 Thioploca ingrica, L40998 Ectothiorhodospira shaposhnikovii, M59151

Discodermia dissoluta, Halichondria okadai, Suberites, Theonella swinhoei (16)

Discodermia dissoluta clone Dd-spF-A85, AY897105 Halichondria okadai clone HOC3, AB054137

Rhopaloeides odorabile (5)

cold marine sediment clone Sva0091, AJ240987 hypersaline lake clone ML1218M-31, AF452606 deep-sea sediment clone BD1-1, AB015514 hydrothermal vent clone MT-NB40, AB186988 Allochromatium vinosum, M26629 Thiocapsa marina, Y12302 Halochromatium salexigens, X98597 Achromatium oxaliferum, AJ010594

Cymbastela concentrica, Homaxinella balfourensis (5)

deep-sea sediment clone MBMPE4, AJ567535 deep-sea sediment clone MBMPE23, AJ567542 deep-sea sediment clone JTB23, AB015248 . deep-sea sediment clone F8, AY375132

Plakortis sp., Theonella swinhoei (3) Sargasso Sea metagenome clone IBEA_CTG_2115585, AACY01032263

Agelas dilatata, Cymbastela concentrica, Plakortis sp., Theonella swinhoei (5)

Aplysina cavernicola, Axinella corrugata, Axinella verrucosa (4)

0.10 192 Suppl. FIG. 8 - Taylor et al.

[Suppl. Fig. 6]

Demospongiae isolate M394, AY371416 Halomonas salina, L42617 Halomonas campisalis, AF054286 Halomonas pantelleriense, X93493 Pachastrellidae isolate N280, AY368510 Erylus formosus isolate CNJ658, AY780448 deep-sea sediment clone wp39, AJ551114 sponge (11 sponges) isolate K354, AY368511 Craniella australiensis DGGE band D5, AY947733 sponge (5 sponges) isolate R261, AY368520 Marinomonas mediterranea, AF063027 Halichondriidae isolate H453, AY371418 bacterioplankton clone Tw-1, AY028196 Suberites clone S2-2, AJ810792 Oleispira antarctica, AJ426420 Isops phlegraei isolate Gp_4_7.1, AM117931 Oceanobacter kriegii, AB006767 Scleritodermidae isolate S018, AY371422 marine clone Osedax_sym1, AY549004 Oceanospirillum beijerinckii, AB006760 Dysidea avara, Halichondria rugosa, Stelletta tenui (6) Stelletta tenui clone 196, DQ236270 Halichondria rugosa clone H.331, DQ274131 Stelletta tenui clone 13, DQ236253 Stelletta tenui clone 117, DQ236266 Stelletta tenui clone 176, DQ236268 Stelletta tenui clone 81, DQ236260 fish-associated clone GA271, DQ263708 Psychrobacter maritimus, AJ609272 "Psychrobacter psychrophilus", AJ748270 Antarctic isolate DVS 9a, AJ577812 Psychrobacter marincola, AJ309941 Psychrobacter submarinus, AJ309940 Arctic sea ice clone ARK10000, AF468382 Craniella australiensis DGGE band D4, AY947732 deep-sea sediment clone wp37, AJ551112 Psychrobacter jeotgali, AF441201 Psychrobacter salsus, AJ539104 Stelletta tenui clone 85, DQ236262 sponge (6 sponges) isolate K512, AY368564 Scleritodermidae isolate K337, AY368565 Psychrobacter pacificensis, AB016057 Antarctic isolate ANT9171, AY167289 Homaxinella balfourensis clone H51, AY321383 Psychrobacter sp. B-5151, DQ399761 sponge (5 sponges & sediment) isolate P672, AY368566 Stelletta tenui clone 87, DQ236263 Stelletta tenui clone 191, DQ236269 Dysidea avara clone D.148, DQ274154 Dysidea avara clone D.21, DQ274135 Dysidea avara clone D.52, DQ274142 Dysidea avara clone D.10, DQ274134 Dysidea avara clone D.84, DQ274145 Dysidea avara clone D.59, DQ274143 Psychrobacter okhotskensi, AB094794 Psychrobacter urativorans, U46141 Psychrobacter glacincola, U46145 Psychrobacter immobilis, U39399 Psychrobacter alimentarius, AY513646 Dysidea avara clone D.142, DQ274153 Rhabdastrella globostellata isolate HRG13, AY372913 Pseudoceratina clavata isolate HPC13, AY372933 Psychrobacter faecalis, AJ421528 Psychrobacter pulmonis, AJ437696 Halichondria rugosa clone H.38, DQ274121 Halichondria rugosa clone H.272, DQ274128 Arctic clone ARKCH4-16 , AF468266 Stelletta tenui DGGE band A4, AY947756 Craniella australiensis DGGE band D7, AY947735 Stelletta tenui DGGE band A2, AY947754 Craniella australiensis DGGE band D2, AY947730 Halichondria rugosa clone H.58, DQ274122 Halichondria rugosa clone H.195, DQ274126 Stelletta tenui clone 94, DQ236265 Stelletta tenui DGGE band A3, AY947755 Stelletta tenui DGGE band A5, AY947757 Stelletta tenui DGGE band A7, AY947759 Stelletta tenui DGGE band A8, AY947760 Stelletta tenui DGGE band A6, AY947758 Stelletta tenui DGGE band A1, AY947753 Stelletta tenui DGGE band A9, AY947761 Moraxella cuniculi, AF005189 Moraxella lacunata, AF005170 Moraxella osloensis, AF005190 activated sludge clone T17, Z93992 deep-sea mud clone HTA608, AB002658 sponge (7 sponges) isolate E929, AY362003 Axinella corrugata clone OS024, AY542770 Stelletta tenui clone A06, DQ274111 Acinetobacter tjernbergiae, AF509825 Halichondria DGGE band B2, AY947722 Halichondria DGGE band B3, AY947723 Halichondria DGGE band B1, AY947721 Antho chartacea clone AnCha211f, EF076235 marine sediment clone NANK-B103, AY436526 Acinetobacter lwoffii, X81665 sponge (6 sponges) isolate H742, AY362004 sponge (Pachastrellidae + coral) isolate J332, AY362002 Acinetobacter calcoaceticus, X81661 Acinetobacter ursingii, AJ275038 , X81660 Cymbastela concentrica (from 2 locations) (5) marine clone A714013, AY907800 Sargasso Sea metagenome clone IBEA_CTG_2032883, AACY01093622 marine bacterioplankton clone AEGEAN_133, AF406527 Rhopaloeides odorabile clone R180, AF333528 marine clone HTCC2188, AY386344 dinoflagellate-associated clone GCDE08_AB, AY701465 Axinella corrugata clone OS040, AY542786 Sargasso Sea metagenome clone IBEA_CTG_2134086, AACY01044896 marine bacterioplankton clone AEGEAN_206, AF406533 Halichondria panicea clone HRV33, Z88589 Phyllospongia papyracea clone 34P16, AY845231 bacterioplankton clone ZA3412c, AF382137 Axinella verrucosa clone E01-9C-12, AJ581349 sponge (6 sponges & coral) isolate K461, AF489287 sponge (6 sponges & coral) isolate K456, AY362013 deep-sea hydrothermal vent clone EPR 6, AY394865 deep-sea sediment clone MBAE17, AJ567578 sponge (6 sponges & coral) isolate N331, AY362014 sponge (2 sponges) isolate D529, AY362015 Alcanivorax venustensis, AF328762 sponge (2 sponges) isolate P653, AY362012 Alcanivorax jadensis, AJ001150 Alcanivorax borkumensis, Y12579 Kirkpatrickia varialosa DGGE band C, AY320214 marine clone Arctic96BD-1, AF354605 Discodermia dissoluta clone Dd-spF-A6, AY897073 seagrass clone HstpL43, AF159674 Halichondria okadai clone HOC27, AB054161 Halichondria okadai clone HOC2, AB054136 Theonella swinhoei clone JAWS16, AF434972 Theonella swinhoei clone JAWS20, AF434976 sponge (Thorectidae & gastropod) isolate H262, AY370006 sponge (2 sponges) isolate H425, AY370008 sponge isolate Spongiobacter nickelotolerans, AB205011 Coelosphaeridae, Mycale laxissima, Pachastrellidae (7) Elysia ornata (sea slug) isolate IAM 15107, AB196667 coral clone OTU 5, DQ312238 coral clone OTU 9, DQ312242 coral clone OTU 7, DQ312240 coral clone PDA-OTU3, AY700601 Rhopaloeides odorabile clone NWCu007, AF313496 Callyspongia sp., Halichondria okadai (6) Zooshikella ganghwensis, AY130994 Haliclona isolate HAL31, AJ849369 Petrosiidae isolate L971, AY371453 Arctic clone FE8, AJ557845 Arctic clone ARKDMS-57, AF468259 Petrosiidae isolate L977, AY371452 sponge (11 sponges & sea urchin) isolate F886, AY368519 Marinobacter excellens, AY180101 Marinobacter aquaeolei, AJ000726 Oleiphilus messinensis, AJ295154 fuel cell clone oc58, AY491599 [Suppl. Figs. 7 & 9]

Homaxinella balfourensis, Kirkpatrickia varialosa, Latrunculia apicalis, Mycale acerata, Sphaerotylus antarcticus (6) Mycale acerata clone M15, AY321410 Stylinos sp. DGGE band 13, AY258611 Suberites clone S15, AJ810788 fish farm sediment clone Nubeena400, AY499935 Chondrilla nucula clone Cnuc42, DQ079043 Axinella verrucosa clone E01-9C-26, AJ581351 Kirkpatrickia varialosa DGGE band A, AY320212 sponge (Kirkpatrickia varialosa, Mycale acerata, Latrunculia apicalis & seawater) DGGE band E, AY320216 salt marsh clone SIMO-2077, AY711443 ML, MP Axinella corrugata, Chondrilla nucula, Microciona sp., Suberites sp. (8)

Axinella corrugata, Homaxinella balfourensis, Kirkpatrickia varialosa, Sphaerotylus antarcticus (10) coral DGGE band 8, DQ312256 coral DGGE band 9 , DQ312257 Kirkpatrickia varialosa DGGE band B, AY320213 marine clone agg47, L10949 marine fosmid clone EF100-93H11, AY627369 Thioalkalispira microaerophila, AF481118 Latrunculia apicalis clone L28, AY321398 Antarctic lake clone WLB16-163, DQ015859 Leucothrix mucor, X87277 Methylobacter luteus, M95657 Methylomonas aurantiaca, X72776 Rhopaloeides odorabile clone R7, AF333541 moth testis clone, L22481 aquatic clone 196up, AY212649 sponge (4 sponges, sediment, polychaete) isolate F802, AY371433 Axinella corrugata clone OS011, AY542757 Stenotrophomonas maltophilia, AB180661 Halichondria DGGE band B8, AY947728 sponge (2 sponges & sea cucumber) isolate F769, AY369998 brewery bottling plant clone MTAA44, AJ619029 Demospongiae isolate M967, AY368563 Pseudoxanthomonas broegbernensis, AJ012231. Xanthomonas campestris, X95917 Pseudoceratina clavata isolate HPC8, AY372931 Xylella fastidiosa, AF536769 Rhopaloeides odorabile clone R33, AF333535 penguin dropping sediment clone KD2-45, AY218584 israelensis, X73408 , M59157 to , Z21931 Piscirickettsia salmonis, U36941 outgroup , M35014

0.10 193 Suupl. FIG. 9 - Taylor et al.

Tedania ignis DGGE band #8, AY337327 Aplysina archeri DGGE band, AY337334 Xestospongia muta DGGE band, AY337336 Chondrilla nucula DGGE band #11, AY337324 Lissodendoryx issodyctialis DGGE band #10, AY337329 Haliclona implexiformis DGGE band #4, AY337325 Geodia papyracea DGGE band #5, AY337326 Geodia papyracea DGGE band #6, AY337323 Tedania ignis DGGE band, AY337322 Nitrosomonas europaea, BX321856 Svenzea zeai DGGE band, AY337335 Nitrosomonas eutropha, AY123795 Nitrosomonas communis, AF272417 Nitrosomonas marina, AF272418 Nitrosomonas oligotropha, AF272422 Nitrosospira briensis, AY123800 Nitrosovibrio tenuis, AY123803 Gallionella ferruginea, L07897 Rhodocyclus tenuis, D16208 Azoarcus tolulyticus, L33691 microbial fuel cell clone oca12, AY491608 Thauera aromatica, X77118 Sterolibacterium denitrificans, AJ306683 uranium-contaminated groundwater clone 015d-B11, AY661971 Aplysina cavernicola DGGE band 9, AY180084 Aplysina cavernicola DGGE band 11, AY180086 Aplysina cavernicola DGGE band 3, AY180078 Aplysina cavernicola DGGE band 2, AY180077 Aplysina cavernicola DGGE band 10, AY180085 Burkholderia cepacia, M22518 Burkholderia stabilis, AF148554 Burkholderia pseudomallei, AY305785 Burkholderia thailandensis, U91838 Burkholderia caledonica, AF215704 Burkholderia fungorum, AF215705 Halichondria panicea clone HNSM13, Z88568 Halichondria panicea clone HNSS13, Z88571 Burkholderia sacchari, AF263278 Spongilla lacustris clone SL52, AY466484 Spongilla lacustris clone SS23, AY598787 Spongilla lacustris clone SL04, AY466483 freshwater clone LD17, Z99998 Polynucleobacter necessarius, X93019 Ralstonia solanacearum, X67036 Chromobacterium violaceum, M22510 , AF310493 Sutterella wadsworthensis, L37786 deep-sea sediment clone BJS72-013, AB238978 Alcaligenes faecalis, M22508 sponge (2 sponges & coral) isolate L981, AY371437 Stelletta tenui clone A11, DQ274113 Stelletta tenui clone A18, DQ274114 Halichondria rugosa clone B81, DQ277989 Dysidea avara DGGE band C2, AY947740 Alcaligenes defragrans, AJ005447 Bordetella bronchiseptica, U04948 , U04950 sponge (2 sponges) isolate N123, AY364595 sponge (6 sponges & other invertebrates) isolate N317, AY362011 Achromobacter piechaudii, AB010841 Acidovorax defluvii, Y18616 Variovorax paradoxus, AJ420329 Comamonas testosteroni, M11224 Halichondria sp. DGGE band B5, AY947725 humic lake clone CrystalBog2KD10, AY792258 Dysidea avara DGGE band C11, AY947749 Leptothrix discophora, Z18533 Thiomonas intermedia, AY455809 Antho chartacea clone AnCha2_16, EF076225 Antho chartacea clone AnCha2_17, EF076226 Antho chartacea clone AnCha2_21, EF076228 Antho chartacea clone AnCha2_7, EF076213 Antho chartacea clone AnCha2_31, EF076223 Antho chartacea clone AnCha2_8, EF076214 Antho chartacea clone AnCha2_18, EF076227 Antho chartacea clone AnCha2_10, EF076215 Antho chartacea clone AnCha2_26, EF076207 Antho chartacea clone AnCha2_15, EF076224 Antho chartacea clone AnCha2_24, EF076210 Antho chartacea clone AnCha2_27, EF076208 Antho chartacea clone AnCha2_5, EF076211 Antho chartacea clone AnCha1_31, EF076220 Antho chartacea clone AnCha1_32, EF076221 Antho chartacea clone AnCha1_19, EF076204 Antho chartacea clone AnCha1_8, EF076198 Antho chartacea clone AnCha1_13, EF076200 Antho chartacea clone AnCha1_15, EF076202 Antho chartacea clone AnCha1_21, EF076205 Antho chartacea clone AnCha111f, EF076241 Antho chartacea clone AnCha1_2, EF076195 Microciona clone M6, AJ810798 Microciona clone M4, AJ810797 Mycale acerata clone M13, AY321407 Mycale acerata clone M140, AY321408 Mycale acerata DGGE band M, AY320224 Stylinos sp. DGGE band 4, AY258602 Stylinos sp. DGGE band 15, AY258613 Latrunculia apicalis clone L39, AY321400 Latrunculia apicalis clone L48, AY321423 Mycale acerata clone M23, AY321412 Latrunculia apicalis clone L44, AY321402 Latrunculia apicalis clone L8, AY321403 Latrunculia apicalis clone L35, AY321419 Latrunculia apicalis clone L9, AY321404 Halichondria panicea clone HRV10, Z88586 Halichondria panicea clone HRV40, Z88592 to Suppl. Kirkpatrickia varialosa DGGE band L, AY320223 Fig. 7 waste gas biofilter clone BIji21, AJ318149 activated sludge clone 15, AF513088 Chondrilla nucula clone Cnuc13, DQ079054 deep-sea sediment clone B488, AY375076

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194 PAPER II

Diversity and mode of transmission of ammonia- oxidizing archaea in marine sponges

Doris Steger,1 Piers Ettinger-Epstein,2 Stephen Whalan,2 Ute Hentschel,3 Rocky de Nys,2 Michael Wagner1 and Michael W. Taylor14

1Department of Microbial Ecology, University of Vienna, Althanstr. 14, A-1090 Vienna, Austria. 2School of Marine and Tropical Biology, James Cook University, Townsville, Qld 4811, Australia. 3Research Center for Infectious Diseases, University of Würzburg, 97070 Würzburg, Germany. 4School of Biological Sciences, University of Auckland, Private Bag 92010, Auckland, New Zealand.

Concept by D.S. and M.W.T. Sampling was done by P.E-E., S.W., U.H. and R.N. D.S. performed all experiments. M.W.T., M.W. and D.S. wrote the manuscript.

Environmental Microbiology (2008) 10(4), 1087–1094 doi:10.1111/j.1462-2920.2007.01515.x

Brief report

Diversity and mode of transmission of ammonia-oxidizing archaea in marine sponges

Doris Steger,1 Piers Ettinger-Epstein,2 eukaryotic microorganisms (Taylor et al., 2007). Sponge- Stephen Whalan,2 Ute Hentschel,3 Rocky de Nys,2 associated archaea were first discovered in 1996, with Michael Wagner1* and Michael W. Taylor1,4** the notable finding of a psychrophilic crenarchaeote, 1Department of Microbial Ecology, University of Vienna, ‘Candidatus Cenarchaeum symbiosum’, within the Althanstr. 14, A-1090 Vienna, Austria. Californian sponge Axinella mexicana (Preston et al., 2School of Marine and Tropical Biology, James Cook 1996). ‘C. symbiosum’ is consistently associated with University, Townsville, Qld 4811, Australia. A. mexicana, comprising up to 65% of prokaryotic cells 3Research Center for Infectious Diseases, University of within the sponge (Preston et al., 1996), and has become Würzburg, 97070 Würzburg, Germany. an important model organism within the field of environ- 4School of Biological Sciences, University of Auckland, mental genomics (Schleper et al., 2005; Hallam et al., Private Bag 92010, Auckland, New Zealand. 2006a,b). Numerous reports of archaea associated with sponges have since emerged (Webster et al., 2001a; Margot et al., 2002; Lee et al., 2003; Holmes and Blanch, Summary 2006; Pape et al., 2006), and it is now apparent that The model marine crenarchaeote ‘Cenarchaeum sym- archaea form part of the natural microbiota of many biosum’ is until now the only ammonia-oxidizing demosponges. archaeon known from a marine sponge. Here, phy- With a handful of reported exceptions (Webster et al., logenetic analyses based on the 16S rRNA and 2001a; Holmes and Blanch, 2006), archaea within marine ammonia monooxygenase subunit A (amoA) genes sponges are members of the phylum Crenarchaeota. The revealed the presence of putative ammonia-oxidizing ubiquity and abundance of crenarchaeotes within marine archaea (AOA) in a diverse range of sponges from the systems is beyond question (DeLong, 1992; Karner et al., western Pacific, Caribbean and Mediterranean. amoA 2001), yet until recently their likely functional role(s) was diversity was limited even between different oceans, unknown. The 2005 isolation in pure culture of an with many of the obtained sequences (75.9%; ammonia-oxidizing marine crenarchaeote (Könneke et al., ntotal = 83) forming a monophyletic, apparently 2005) confirmed earlier predictions from environmental sponge- (and coral-) specific lineage, analogous to genomics studies (Venter et al., 2004; Treusch et al., those previously inferred from comparative 16S rRNA 2005) that crenarchaeotes could be capable of chem- gene studies of sponge-associated microbes. The olithoautotrophic ammonia oxidation. Subsequent recov- presence of AOA in sponge larvae, as detected by 16S ery of archaeal ammonia monooxygenase (amoA) gene rRNA and amoA PCR assays as well as by fluores- sequences from varied seawater and sediment samples, cence in situ hybridization, suggests they are verti- together with compelling abundance and process data, cally transmitted and thus might be of importance for indicates that archaeal ammonia oxidation may be wide- ammonia detoxification within the sponge. spread and important in marine systems (Francis et al., 2005; Ingalls et al., 2006; Wuchter et al., 2006; Coolen et al., 2007; Lam et al., 2007; Mincer et al., 2007). Introduction Given the frequent association of Crenarchaeota with marine sponges, together with evidence of nitrification Many marine sponges harbour diverse and abundant within these host organisms (Corredor et al., 1988; microbial communities, including bacteria, archaea and Diaz and Ward, 1997; Bayer et al., 2007), it seems likely Received 19 July, 2007; accepted 27 October, 2007. For that ammonia-oxidizing archaea (AOA) should occur in correspondence. *E-mail [email protected]; Tel. (+43) sponges. Indeed, Hallam and coworkers recently 1 4277 54390; Fax (+43) 1 4277 54389; **E-mail mw.taylor@ auckland.ac.nz; Tel. (+64) 9 3737599 ext 82280; Fax (+64) reported the presence of amo gene homologues within 9 3737416. the sequenced genome of the sponge symbiont

197 Part I

Table 1. Details of sponge samples used for this study.

Sponge species Order/Family Collection location (latitude, longitude)

Axinella polypoides Halichondrida/Axinellidae Rovinj, Croatia (45°05′N, 13°38′E) Chondrosia reniformis Chondrosida/Chondrillidae Rovinj, Croatia (45°05′N, 13°38′E) Aplysina insularis Verongida/Aplysinidae Little San Salvador Island, Bahamas (24°35′N, 75°58′W) Cliona sp. Hadromerida/Clionaidae Little San Salvador Island, Bahamas (24°35′N, 75°58′W) Plakortis sp. Homosclerophorida/Plakinidae Little San Salvador Island, Bahamas (24°35′N, 75°58′W) Coscinoderma sp. Dictyoceratida/Spongiidae Orpheus Island, Queensland, Australia (18°36′S, 146°29′E) Luffariella variabilis Dictyoceratida/Thorectidae Orpheus Island, Queensland, Australia (18°36′S, 146°29′E) Rhopaloeides odorabile Dictyoceratida/Spongiidae Pelorus Island, Queensland, Australia (18°34′S, 146°29′E) Callyspongia sp. Haplosclerida/Callyspongiidae Bare Island, Sydney, Australia (33°59′S, 151°14′E) Siphonochalina sp. Haplosclerida/Callyspongiidae Bare Island, Sydney, Australia (33°59′S, 151°14′E) Stylinos sp. Halichondrida/Halichondriidae Bare Island, Sydney, Australia (33°59′S, 151°14′E)

In all cases, three individuals of each sponge species were collected. All listed sponges are in the class Demospongiae.

‘C. symbiosum’ (Hallam et al., 2006a,b). Also present in Aplysina insularis, Cliona sp. and Plakortis sp., the Medi- the ‘C. symbiosum’ genome are other genes encoding terranean sponges Axinella polypoides and Chondrosia components associated with the ammonia oxidation reniformis, the Great Barrier Reef sponges Coscinoderma pathway, including urease and nitrite reductase. Intrigu- sp., Luffariella variabilis and Rhopaloeides odorabile and ingly, no homologue was found for hydroxylamine the south-eastern Australian sponges Callyspongia sp., oxidoreductase (hao), a key component of ammonia Siphonochalina sp. and Stylinos sp. (Table 1). All of these oxidation in bacteria (Hallam et al., 2006a). sponges are abundant at their respective collection Confirmation of the occurrence of AOA within a marine locations. There was no clear biogeographic effect, with sponge brings with it many questions. For example, are many of the sponge-derived sequences forming a large AOA associated with many sponges, or merely limited to monophyletic cluster comprising sequences from the the ‘C. symbiosum’–A. mexicana symbiosis? What is the Great Barrier Reef, the Bahamas, the Mediterranean and magnitude of AOA diversity within sponges, that is do California (= ‘C. symbiosum’) (Fig. 1). Also present in this different phylotypes occur or are they all phylogenetically cluster are several coral-derived crenarchaeotal amoA closely affiliated with ‘C. symbiosum’? How are AOA sequences (Beman et al., 2007). Although the relatively transferred between generations of the host sponge? limited archaeal amoA data set (n = 1547 in the database Addressing such questions is important in order to provide we used) currently available constrains speculation, this a sound framework for subsequent, detailed studies of putatively ‘sponge- and coral-specific’ crenarchaeotal nitrification in sponges. The accumulation of metabolic cluster is reminiscent of those reported previously for the wastes (including ammonia and nitrite) may have adverse 16S rRNA gene (Taylor et al., 2007; see also Fig. 2). It will effects on sponge health, with evidence for the detrimen- be of interest to see whether the cluster remains intact tal impact of nitrite on juvenile sponge development once more marine amoA data are obtained, but it is note- (Kahlert and Neumann, 1997). The ability to metabolize worthy that none of the more than 920 amoA sequences nitrogenous waste products may thus be the basis for a from marine sediments and water samples, including 45 mutualistic relationship between ammonia and nitrite oxi- amoA sequences deposited in the marine metagenomic dizers and marine sponges, and it is important to improve data sets (Venter et al., 2004; Rusch et al., 2007; our understanding of the basic ecological features of the Seshadri et al., 2007), are members of this cluster. Other sponge–AOA association. sponge-derived amoA sequences obtained in this study were similar to those from other marine environments. Several sponges contained more than one amoA Results and discussion sequence type (Fig. 1). Overall, amoA variability in To establish whether AOA are frequent associates of sponges was relatively low (lowest similarity value of two marine sponges, we screened sponges sampled from the retrieved sponge-derived amoA sequences was 94.7% Caribbean, Mediterranean and western Pacific, for the amino acid identity), even among sequences obtained presence of archaeal amoA genes. Although possession from different oceans, although it cannot be ruled out that of this gene does not provide unequivocal proof that a further sequencing may have uncovered a greater diver- given organism can oxidize ammonia, it is strongly sug- sity of phylotypes. The recognized amoA diversity of gestive and we can label such archaea as putative sponges is lower than that of corals and marine sedi- ammonia oxidizers. amoA sequences were obtained from ments, while a comparison to the AOA amoA diversity each of three tested individuals of the Caribbean sponges retrieved from the marine water column revealed a more

198 Microorganisms in marine sponges

GBR Luffariella variabilis adults and larvae clones nA=15 nL=2 Cliona sp. clone n=3 Bah Luffariella variabilis adults and larvae clones nA=1 nL=5 GBR Luffariella variabilis larva clone LuffL19 GBR EF382621, coral clone PA6-27 Aplysina insularis clone 25_3 Bah Plakortis sp. clone n=5 Bah Aplysina insularis clone n=2 Bah EF382505, coral clone MA7-6 Axinella polypoides clone n=3 Med 630004020, “Cenarchaeum symbiosum”

Luffariella variabilis larvae clones nL=3 GBR Rhopaloeides odorabile adults and larvae clones nA=13 nL=7 GBR Chondrosia reniformis clone n=2 Med

EF382451, coral clone DS4-15 Coscinoderma sp. clone Cosc9 GBR EU025171, sediment clone S10-A-14 EF382609, coral clone PA6-15 Luffariella variabilis adult clone LuffA10 GBR EF382501, coral clone MA7-2 Coscinoderma sp. clone n=2 GBR EF382512, coral clone MA7-13 Cliona sp. clone n=2 Bah EF382620, coral clone PA6-26 ”Nitrosopumilus maritimus” and related sequences DQ333407, sea water clone HF770_C01 Siphonochalina sp. clone Siph_2 Syd EF106927, subtropical gyre clone HF770_12N24 Siphonochalina sp. clone Siph_3 Syd DQ148825, Monterey Bay clone MB_C130m_7 DQ148689, Black Sea gyre clone BS15.7_11 EF414241, Black Sea water clone BS2-130mC2 Stylinos sp. clone Styl_2 Syd DQ148846, Monterey Bay clone MB_M2_40m_6 Stylinos sp. clone Styl_1 Syd Stylinos sp. clone Styl_3 Syd Siphonochalina sp. clone Sipho3 DQ148842, Monterey Bay clone MB_M2_40m_2 DQ148766, north Pacific clone ETNP_23 EF106932, subtropical gyre clone HF130_29B4 Siphonochalina sp. clone Siph_1 Syd EU025169, sediment clone S10-A-12 Callyspongia sp. clone Callmp3 Syd EF106943, subtropical gyre clone HF200_15E20 Siphonochalina sp. clone n=2 Syd Callyspongia sp. clone n=3 Syd

Group I.1b

0.10

Fig. 1. amoA-based phylogenetic consensus tree of AOA affiliated with the marine group I.1a Crenarchaeota including sponge-derived sequences (in bold). Polytomies indicate that respective branching order could not be unambiguously resolved by different treeing methods. Codes in shaded boxes indicate collection location: Bah = Bahamas, GBR = Great Barrier Reef, Med = Mediterranean, Syd = Sydney.

The large box represents a monophyletic cluster consisting exclusively of sponge- and coral-derived sequences. nL = number of identical sequences obtained from the respective larvae; nA = number of identical sequences obtained from adult sponges. Arrows indicate the two known ammonia-oxidizing archaea, ‘Candidatus Cenarchaeum symbiosum’ and ‘Cand. Nitrosopumilus maritimus’. Filled circles indicate parsimony bootstrap values (100 resamplings) of 90–100%. Bar, 10% sequence divergence. For sequences obtained during this study, DNA extractions were performed on 5 mg of freeze-dried sponge tissue using the UltraClean Soil DNA Kit (MoBio Laboratories, USA), followed by PCR modified slightly from the protocol of Francis and colleagues (Francis et al., 2005). The cycling conditions were: 94°C for 3 min; 35 cycles of 94°C for 40 s, 53°C for 40 s, 72°C for 40 s; 72°C for 20 min. Cloning was carried out using the TopoTA Cloning Kit (Invitrogen, USA). The displayed consensus tree was constructed based on distance (Fitch), maximum parsimony and maximum likelihood phylogenetic analyses conducted in ARB (Ludwig et al., 2004), with 585 characters used for tree construction. The tree was calculated with a larger data set than displayed, but to improve clarity those clusters (with the exception of the ‘N. maritimus’ cluster) which did not contain sponge-derived sequences were removed after calculation. complicated picture (Fig. 3). More amoA operational taxo- > 90%) the OTU number of planktonic AOA by far exceeds nomic units (OTUs) are found in sponges than in marine the one observed in sponges. For ammonia-oxidizing water samples, if a nucleic acid sequence similarity bacteria 80% sequence similarity of the amoA gene can threshold of 80–85% is applied, while at the microdiversity be used as a threshold for species differentiation level (sequence similarity threshold for OTU definition (Purkhold et al., 2000). Under the assumption that this

199 Part I

Rhopaloeides odorabile adults and larvae clone group nL=33 nA=36 AY029297, Rhopaloeides odorabile clone Arc2 AY192628, Halichondria sp. clone Ar 12-1 AY372934, Rhabdastrella globostellata clone DirRG DQ424909, Astrosclera willeyana clone A11B2 DQ424910, Stelligera stuposa clone A11B1 DQ299290, Didiscus sp. clone A11B4 DQ299268, Geodia media clone A10A11 AY192645, Petrosia sp. clone Ar24-5 DQ299270, Scopalina reutzleri clone A307a DQ299264, Petrosia sp. clone A253A AB019725, hydrothermal vent clone pMC1A103 DQ299267, Reinochalina stalagmites clone A5E7 DQ085103, marine aquarium clone SeAq_T2A1 DQ085097, “Nitrosopumilus maritimus“ DQ299273, Phakellia sp. clone A15D3 DQ641842, Madovi Estuary sediment clone MES-137 DQ299271, Acanthella sp. clone ArB12 DQ299272, Axinella polypoides clone A9A3 DQ299263, Astrosclera willeyana clone A11B3 Plakortis sp. clone PKA26 AY192627, Spirastrella panis clone Ar08-1 DQ299262, Reinochalina stalagmites clone A5D8 AF419636, hydrothermal sediment clone C1_R010 DQ363813, mangrove soil clone MKCST-dx11 AJ347774, brine-seawater clone ST-3K4A AY192630, Sarcotragus sp. clone Ar19-17 U71114, marine clone C20 U71112, marine clone C6 AB193986, hydrothermal vent clone Sc-EA02 AY320198, Mycale acerata clone 1 AF119126, deep sea sediment clone CRA7-11cm AF119130, deep sea sediment clone CRA20-0cm AJ347775, marine clone ST-12K13A AY320200, Mycale acerata clone 3 AF419646, hydrothermal sediment clone CS_R008 AB019723, hydrothermal vent clone pMC2A1 AY396000, beach microcosm sediment clone BC-A AY320199, Mycale acerata clone 2 AB019722, deep-sea hydrothermal vent clone pIVWA101 AF393466, marine fosmid clone 74A4 AY320201, Mycale acerata clone 4 AY192644, Halichondria okadai clone Ar18-1 Luffariella sp. adult clone LA_7 DQ085101, marine aquarium clone SeAqRB01 DQ299265, Suberites sp. clone A17A AY192633, Erylus nobilis clone Ar25-51 DQ299266, Axinella polypoides clone app654 AY192639, Sarcotragus sp. clone Ar19-24 AY192634, Halichondria sp. clone Ar12-16 AY192643, Petrosia sp. clone Ar24-16 AY192642, sponge clone Ar23-28 AY192632, sponge clone Ar23-13 AY192640, Sarcotragus sp. clone Ar19-36 AY192641, sponge clone Ar23-23 AB050232, gold mine clone SAGMA1 AY082452, acid mine clone 44a-A1-1 AF119138, deep-sea sediment clone APA4-0cm DQ299274, Ptilocaulis sp. clone A13F

Luffariella sp. adults and larvae clone group 1 nL=11 nA=16

Luffariella sp. adults and larvae clone group 2 nL=1 nA=20 AF186423, Theonella swinhoei clone PAAR8

Luffariella sp. adults and larvae clone group 3 nL=38 nA=49 AY592447, deep-sea sediment clone Napoli-1A-14 DQ424908, Axinella corrugata clone BHF4 DQ299280, Prosuberites laughlini clone A10A3 AF421159, Axinella sp. clone AF420236, Axinella damicornis clone AF420237, Axinella verrucosa clone DQ299278, Aciculites sp. clone A10A5 DQ417470, sea floor clone a87R34 AF119134, deep-sea sediment clone APA1-0cm AY505049, hydrothermal chimney clone LC1231a76 AY192629, Halichondria okadai clone Ar18-3 DQ299277, Axenella polycapella clone A7G11 DQ299276, Axinella aurensis clone A165A AB193956, hydrothermal vent clone MaSc-NEA01 AB019727, hydrothermal vent clone pIVWA1 U78199, marine clone SB95-57 U71109, marine bacterioplankton clone PM7 DQ085102, marine aquarium clone SeAqRB09 AF119127, deep-sea sediment clone CRA8-11cm Agelas dilatata clone ADA9 DQ299275, Axinella corrugata clone A137.2 AY192631, sponge clone Ar20-3 U51469, Axinella mexicana clone “Cenarchaeum symbiosum“

Luffariella sp. larvae clone group 4 nL=6 DQ299279, Axinella polypoides clone A9A4 SAGMCG1 AB050228, gold mine clone SAGMA-W AJ535122, mining waste clone Gitt-GR-74 AF227640, plant root clone TRC23-10 U62816, soil clone SCA1166 AF227644, plant root clone TRC23-38 U62819, soil clone SCA1175 AY192637, Halichondria okadai clone Ar-7

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200 Microorganisms in marine sponges

Fig. 2. 16S rRNA gene-based phylogenetic consensus tree displaying the affiliation of sponge-associated Crenarchaeota within Marine Group I. Shaded boxes represent sponge-specific monophyletic clusters sensu Hentschel and colleagues (Hentschel et al., 2002). Other boxes signify Rhopaloeides odorabile or Luffariella variabilis sequences, with nL = number of identical sequences obtained from the respective larvae and nA = number of identical sequences obtained from adult sponges. Arrows indicate the two known ammonia-oxidizing archaea, ‘Candidatus Cenarchaeum symbiosum’ and ‘Cand. Nitrosopumilus maritimus’. Filled circles indicate parsimony bootstrap values of 90–100%. Bar, 10% sequence divergence. The 16S rRNA gene was amplified using the Arch21F/Arch958R primers (DeLong, 1992) with the following PCR cycling conditions: 94°C for 3 min; 35 cycles of 94°C for 40 s, 55°C for 40 s, 72°C for 1 min; 72°C for 20 min. 879 characters were used for tree construction.

rule can also be applied for AOA, these findings indicate Elucidating the mechanism(s) by which symbionts are that at the species level more AOA occur in sponges than acquired by a host is an important goal in symbiosis in the marine water column (note that 302 marine water research. Numerous microscopy- and molecular-based column sequences and only 83 sponge-related amoA studies have demonstrated transmittance of diverse sequences are available; Fig. 3b), while in the latter envi- microbial assemblages in sponge reproductive stages ronment there is apparently an enormous strain-level (eggs, sperm, larvae) (Usher et al., 2001; 2005; Eresk- diversity of a few species. ovsky et al., 2005; Enticknap et al., 2006; Schmitt et al.,

600 Fig. 3. Archaeal amoA diversity in various 100 environments. A A. The number of amoA operational taxonomic units (OTUs) is plotted against the 500 80 evolutionary distance threshold used for OTU definition. The inset displays a selected region of the graph to enhance clarity. 400 60 B. Rarefaction analysis of the amoA sequences using an evolutionary distance threshold of 20% (i.e. 80% similarity) for OTU 40 300 definition. The analysis was performed using DOTUR (Schloss and Handelsman, 2005).

20 Symbols represent the following environments: ( ) sponge, ( ) coral, ( ) 200 water column, ( ) thermal environments, ( )

no. of OTUs observed 0 soil, ( ) activated sludge, ( ) sediment. The 0.06 0.08 0.10 0.12 0.14 0.16 0.18 0.20 0.22 0.24 0.26 45 archaeal amoA sequences from the 100 metagenome projects were excluded from these analyses owing to a relatively low sequence quality which would create microdiversity artefacts. 0 0.0 0.1 0.2 0.3 0.4 0.5 evolutionary distance

20 B 18 16

14

12

10

8

6 no. of OTUs observed 4

2

0 0 100 200 300 400 500 600 700 no. of amoA sequences

201 Part I

Fig. 4. FISH visualization of Crenarchaeota in ABLuffariella variabilis larvae. A. Blue – Archaea detected by Cy5-labelled Arch915 probe (Stahl and Amann, 1991). Red – Bacteria [Cy3-labelled with the probe set EUB338(I–III) (Daims et al., 1999)]. Green – background fluorescence from sponge larva. B. Blue – Archaea (Cy5-labelled with probe Arch915). Red – Crenarchaeota [Cy3-labelled with probe CREN512 (Jurgens et al., 2000)]. Note that pink signal indicates cohybridization of Arch915 and CREN512 probes. Green – background fluorescence from larva.

2007; Sharp et al., 2007). In the tropical sponge Corticium which had been embedded in Neg-50 (Microm, Richard- sp., this assemblage includes archaea (Sharp et al., Allan Scientific) and frozen, we were able to visualize 2007). However, to our knowledge there exists no infor- Bacteria and Crenarchaeota in larvae from L. variabilis. mation as to whether AOA are transmitted via the repro- Most of these microorganisms were located in the central ductive stages. To address this issue, we collected both part of the larvae (Fig. 4). However, because of unexpect- adult and larval samples from two viviparous Great Barrier edly rapid deterioration of the fixed larvae samples at Reef sponges, L. variabilis (Ettinger-Epstein et al., 2007) -20°C we were unable to perform quantitative FISH or to and Rhopaloeides odorabile (Whalan et al., 2007). The use FISH for either the study of R. odorabile larvae, or for latter species harbours a dense and diverse microbial the application of newly designed probes targeting spe- community including archaea (Webster et al., 2001a,b), cific sequence types in L. variabilis. Unlike many marine whereas nothing is known about the microbiology of invertebrates, the larvae of sponges are lecithotrophic, or L. variabilis. Larvae and corresponding individual parent non-feeding (Maldonado, 2004; Mariani et al., 2006). sponges were sampled during the 2005–2006 austral Sponge larvae thus rely upon finite stored nutrients for summer, with larvae collected by placing larval traps over nourishment, and there exists no evidence that they are each individual sponge following the method of Lindquist capable of feeding upon microorganisms from the sur- and colleagues (1997). Larvae were collected and either rounding seawater. We are therefore confident that larval frozen (for DNA analyses), or preserved in 3% paraform- transmission of AOA is indeed occurring, and that it is not aldehyde or 96% ethanol [for fluorescence in situ hybrid- merely diet-derived archaea that were detected in the ization (FISH); detailed in Daims et al. 2005] within 2 h of larvae by FISH. release. Taken together, our data suggest that AOA may be We PCR-amplified the archaeal amoA and 16S rRNA frequently associated with marine sponges. Their appar- genes from DNA extracts of freeze-dried adult and larval ent transmission in the larval stages of at least one host samples of both sponge species. Phylogenetic analyses species is suggestive of a possible symbiotic role for revealed the same crenarchaeotal 16S rRNA phylotype these microorganisms. Further work is needed to deter- [previously reported from this sponge (Webster et al., mine the contribution of AOA to the nitrification process 2001a)] in both adults and larvae of R. odorabile (Fig. 2). within sponges, and to establish whether – as shown for Similarly, a single, identical amoA sequence type was some other environments (Leininger et al., 2006; Wuchter recovered from R. odorabile adults and corresponding et al., 2006) – archaea and not bacteria are the numeri- larvae (Fig. 1). Both amoA- and 16S rRNA-based analy- cally dominant ammonia oxidizers in marine sponges. ses indicated a more diverse AOA community in L. variabilis (Figs 1 and 2). Again, in most cases the same Acknowledgements sequence types were found in adults and larvae, suggest- ing that AOA are present in the larvae of these two We thank K. Bergauer, R. Radax and K. Stoecker for their sponges. To confirm that archaea were present within the help in different stages of this research. D.S. received a DOC-FORTE stipend from the Austrian Academy of larvae, and did not represent surface contaminants from Sciences. Sponge research at James Cook University was the surrounding seawater, we performed FISH with supported by the Research Advancement Program. The archaeal and crenarchaeal 16S rRNA-targeted oligo- research of UH was supported by SFB567 grant (TPC3) of nucleotide probes. By examining 6 mm sections of larvae the Deutsche Forschungsgemeinschaft. The research of

202 Microorganisms in marine sponges

MWT was funded in part by a grant of the German Federal oxidizing archaea in water columns and sediments of the Ministry for Education and Research (Biolog II) to M.W. The ocean. Proc Natl Acad Sci USA 102: 14683–14688. author M.W. would like to thank the University of Vienna Hallam, S.J., Mincer, T.J., Schleper, C., Preston, C.M., for financial support in the framework of the faculty focus Roberts, K., Richardson, P.M., and DeLong, E.F. (2006a) ‘Symbiosis’ and the university research focus ‘Symbiosis Pathways of carbon assimilation and ammonia oxidation Research and Molecular Principles of Recognition’. suggested by environmental genomic analyses of marine Sequence data from this study have been submitted to the Crenarchaeota. PLoS Biol. 4: e95. DDBJ/EMBL/GenBank databases under the accession Hallam, S.J., Konstantinidis, K.T., Putnam, N., Schleper, C., numbers EU049815–EU049846. Watanabe, Y., Sugahara, J., et al. 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204

Part IV

Summary

Summary

Summary

In the present thesis, functional groups of microorganisms were examined in two extremely different ecosystems known for their highly complex microbial community structure.

The first part of this work deals with the abundance, community dynamics, and biogeography of dsrAB-carrying microorganisms in terrestrial wetlands. The dissimilatory (bi-) sulfite reductase (Dsr) is present in microorganisms that reduce sulfate, sulfite or some organosulfonates or in anaerobic syntrophs. Microorganisms with phylogenetically novel variants of the dissimilatory (bi-)sulfite reductase genes dsrAB were identified earlier in the Schlöppnerbrunnen fen (Lehstenbach catchment, Germany). Using newly developed molecular tools to analyze 16S rRNA and dsrAB gene sequences, a temporally stable but spatially variable composition of the putative sulfate reducing populations could be detected in the highly diverse and Acidobacteria dominated fen. Thereby, the proportion of yet uncharacterized deep-branching dsrAB sequences outnumbered the known syntrophic dsrAB lineages. Furthermore, denaturing gradient gel electrophoresis (DGGE) and clone library-based analyses of eight additional wetlands confirmed the widespread occurrence of these deep-branching dsrAB lineages in peatlands. The dsrB-DGGE pattern was not correlated with geographic distance, but observed variability could be largely explained by measured environmental conditions.

In the second part of the thesis, the use of DNA microarray based methods uncovered significant variations in detected signal intensities of these nucleic acid based hybridization assays. DNA microarrays are regularly used in science as well as in clinical diagnosis for high- throughput multiplex analyte detection. By a simplified array format, systematically arranged spatial gradients were detected, which halved the number of detected targets from the edge to the center of the assay. Additionally, experimental data were confirmed by simulation of passive mass transport indicating diffusion limitations of the target molecules as causing mechanism.

In the last part of this work the microbial communities inhabiting marine sponges were investigated. A comprehensive analysis of all publicly available sponge derived 16S rRNA sequences and further 184 sequences of three hitherto unstudied sponges confirmed the presence of monophyletic, sponge-specific 16S rRNA sequence clusters, with approximately one-third of all sponge-derived sequences falling into such clusters. Furthermore, phylogenetic

207 Part IV analyses based on 16S rRNA and ammonia monooxygenase genes and fluorescence in situ hybridization revealed the presence of putative ammonia-oxidizing archaea (AOA) in a diverse range of sponges from the western Pacific, Caribbean and Mediterranean. The presence of AOA in sponge larvae suggests they are vertically transmitted and thus might be of importance for ammonia detoxification within the sponge.

208 Zusammenfassung

Zusammenfassung

In der vorliegenden Arbeit wurden funktionelle Gruppen von Mikroorganismen in zwei äußerst unterschiedlichen Ökosystemen, die beide eine komplexe mikrobielle Gemeinschaftsstruktur aufweisen, untersucht.

Der erste Teil dieser Arbeit befasst sich mit der Häufigkeit, Populationsdynamik und Verbreitung von dsrAB-tragenden Mikroorganismen in terrestrischen Feuchtgebieten. Mikroorganismen, die Sulfat, Sulfit oder Organosulfonate reduzieren beziehungsweise in syntrophen Gemeinschaften leben, können eine dissimilatorische (Bi-) Sulfit-Reduktase (Dsr) besitzen. Neue, tiefzweigende dsrAB-Sequenzen wurden bereits in früheren Studien im Untersuchungsgebiet Schlöppnerbrunnen (Bayern, Deutschland) nachgewiesen. Unter Verwendung verschiedener molekularbiologischer Techniken zur Analyse von 16S-rRNS- und dsrAB-Gensequenzen, konnte in diesem hoch diversen und von Acidobacteria dominierten Moor eine zeitlich stabile, räumlich jedoch variable Zusammensetzung der potentiell sulfatreduzierenden Populationen gezeigt werden. Hierbei waren die bisher nicht charakterisierten, tiefzweigenden dsrAB- Sequenzen signifikant häufiger als die bekannten syntrophen DsrAB-Linien. Das verbreitete Vorkommen dieser tiefzweigenden DsrAB-Linien wurde mittels dsrB basierter denaturierender Gradientengelelektrophorese (DGGE) in acht weiteren Feuchtgebieten bestätigt. Hierbei konnte das detektierte dsrB-DGGE Bandenmuster nicht durch die geographische Distanz der Feuchtgebiete, sondern durch die vorherrschenden Umweltbedingungen erklärt werden.

Im zweiten Teil dieser Arbeit deckte die Anwendung von DNS-Mikroarray basierten Methoden signifikante Schwankungen der detektierten Fluoreszenzsignale in diesen nukleinsäurebasierten Hybridisierungsassays auf. DNS-Mikroarrays werden regelmäßig in der Wissenschaft sowie zur klinischen Diagnose als Hochdurchsatzanalyse komplexer Proben verwendet. Ein vereinfachtes Arraylayout ermöglichte den Nachweis von systematisch angeordneten räumlichen Gradienten in der Signalintensität, wobei die Anzahl an detektierten Zielmolekülen vom Rand zur Mitte des Mikroarrays halbieren wurde. Modellsimulationen wiesen auf Limitierungen im Massetransport der Zielmoleküle als mögliche Ursache hin.

Im letzten Teil dieser Arbeit wurden die mikrobiellen Gemeinschaften in marinen Schwämmen untersucht. Eine umfassende Analyse aller öffentlich verfügbaren 16S-rRNS-Sequenzen aus

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Schwämmen und weitere 184 Sequenzen von drei bisher nicht untersuchten Schwämmen, bestätigte das Vorhandensein monophyletischer, Schwamm-spezifischer 16S-rRNS Gensequenz-Cluster, wobei etwa ein Drittel der analysierten Sequenzen zu diesen Clustern zugeordnet werden konnte. Darüber hinaus zeigten 16S-rRNS- und Ammonium Monooxygenase-Gen als auch Fluoreszenz in situ Hybridisierung basierte Analysen, die Anwesenheit von potentiell ammoniumoxidierenden Archaea (AOA) in marinen Schwämmen aus dem westlichen Pazifik, der Karibik und des Mittelmeers. Das zusätzliche Vorhandensein dieser AOAs in den entsprechenden Schwamm-Larven weist darauf hin, dass AOAs in Schwämmen vertikal übertragen werden.

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APPENDIX

Publications

Sebastian Lücker, Doris Steger, Kasper Urup Kjeldsen, Barbara J. MacGregor, Michael Wagner and Alexander Loy. 2007. Improved 16S rRNA-targeted probe set for analysis of sulfate-reducing bacteria by fluorescence in situ hybridization. J. Microbiol. Methods. 69: 523-528.

Michael Taylor, Regina Radax, Doris Steger and Michael Wagner. 2007. Sponge- Associated Microorganisms: Evolution, Ecology, and Biotechnological Potential. Microbiol. Mol. Biol. Rev. 71: 295-347

Doris Steger, Piers Ettinger-Epstein, Stephen Whalan, Ute Hentschel, Rocky de Nys, Michael Wagner and Michael Taylor. 2008. Diversity and mode of transmission of ammonia-oxidizing archaea in marine sponges. Environ. Microbiol. 10: 1087-1094.

Alexander Loy, Michael Pester and Doris Steger. Phylogenetic Microarrays for Cultivation-Independent Identification and Metabolic Characterization of Microorganisms in Complex Samples. Book chapter in PCR Mutation Detection Protocols, Springer-Verlag. In review.

Doris Steger, Cecilia Wentrup, Michael Wagner and Alexander Loy. Dynamics of an unusual sulfate reducing population within a highly diverse microbial fen community. To be submitted.

Doris Steger, Roman Stocker, Michael Wagner und Alexander Loy. Spatial gradients govern nucleic acid surface hybridizations. In preparation.

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Oral presentations

Doris Steger, Michael Wagner and Alexander Loy. Diversity of apsA genes in a shallow hydrothermal vent system. 5th International meeting on Novel Techniques In Microbial Ecology, Lunz am See, Austria, June 2005

Doris Steger, Cecilia Wentrup, Manuel Hofer, Michael Wagner and Alexander Loy. RING-FISH and qPCR in Schlöppnerbrunnen samples. 6th International workshop on New Techniques In Microbial Ecology, Sæby, Denmark, October 2007

Doris Steger, Cecilia Wentrup, Manuel Hofer, Michael Pester, Michael Wagner and Alexander Loy. New insights in the sulfate reducing community of an acidic fen system. VAAM Jahrestagung, Frankfurt, Germany, March 2008

Poster presentations

Doris Steger, Stephanie Füreder, Stephan Duller, Michael Wagner, Monika Bright and Alexander Loy. Shallow Water Hydrothermal Vents at the Island of Dominica (Caribbean Sea) are Diversity Hot Spots of Sulfur Prokaryotes. ChEss workshop: Atlantic Equatorial Belt, Barcelona, Spain, March 2006

Michael Taylor, Doris Steger, Regina Radax and Michael Wagner. Marine benthic eukaryotes as reservoirs of microbial diversity. International Census of Marine Microbes, Noordwijkerhout, the Netherlands, June 2006

Doris Steger, Regina Radax, Steven Whalan, Piers Ettinger-Epstein, Rocky de Nys, Michael Wagner and Michael Taylor. Ammonia-oxidizing archaea are widespread in marine sponges. 11th International symposium on Microbial Ecology, Vienna, Austria, August 2006.

Regina Radax, Doris Steger, Michael Wagner and Michael Taylor. Phylogeny of all known sponge-associated prokaryotes: do sponge-specific clusters really exist? 11th International symposium on Microbial Ecology, Vienna, Austria, August 2006.

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Curriculum vitae

Name: Doris Steger Date of birth: 27.10.1979 Place of birth: Bruneck, Italy Citizenship: Italy

Education:

1985-1990 Primary school, Pfalzen, Italy 1990-1993 Secondary school, Bruneck, Italy 1993-1998 High school of natural sciences, Bruneck, Italy

1998-1999 Studies of Psychology and nutritional science at the University of Vienna, Austria 1999-2004 Studies of Biology at the University of Vienna, Austria Main subjects: Microbiology and Genetics 2003 Study visit as an Erasmus-student at the Universitá degli Studi di Firenze (Italy) 2004-2005 Diploma Thesis at the Department of Microbial Ecology, Faculty of Life Sciences, University of Vienna, Austria, with the title: “Phylogenie der schwefeloxidierenden Prokaryoten und Analyse ihrer Diversität in einem marinen Flachwasserhydrothermalsystem.” 2005-2009 Ph.D studies at the Department of Microbial Ecology, Faculty of Life Sciences, University of Vienna, Austria, supervised by Prof. Dr. Michael Wagner

since 2010 Clinical Safety manager, Sandoz GmbH, Kundl, Austria

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Acknowledgments

Während der Zeit meiner Doktorarbeit haben mich sehr viele Menschen in verschiedenster Art und Weise unterstützt. An dieser Stelle gilt mein herzlichster Dank:

Michael Wagner, für die Möglichkeit meine Doktorarbeit in seiner Arbeitsgruppe zu erstellen, seine Begeisterung für die Wissenschaft und für die vielen Anregungen, welche mir immer wieder den nötigen Antrieb gegeben haben

Alexander Loy, für seine finanzielle und fachliche Unterstützung, sowie für seine wertvollen Ratschläge bei der Durchführung dieser Arbeit

Michael Taylor, der mit seinem Optimismus, Fachwissen und vielen Ideen sehr zum erfolgreichen Weiterkommen in der ersten Phase meiner Doktorarbeit beigetragen hat

Dem ganzen DOME-Team, für das nette Arbeitsklima, für alle guten Gespräche und für die Hilfsbereitschaft bei der Durchführung der Arbeit. Besonders bedanken möchte ich mich hier bei meinen Arbeitskollegen Steffi, Susi, Kilian, Micha, Baroni, Schmitzi, Basti und Elena, sowie meinen zahllosen Diplomanden und Praktikanten – für die Motivation und fachliche Unterstützung, aber auch für viel Spaß, Verständnis und kurze Weile.

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